Soil Enzymology (Soil Biology, Volume 22)

Soil Biology Volume 22

Series Editor Ajit Varma, Amity Institute of Microbial Technology, Amity University Uttar Pradesh, Noida, UP, India For further volumes: http://www.springer.com/series/5138

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Girish Shukla

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Ajit Varma

Editors

Soil Enzymology

Editors Dr. Girish Shukla Biological, Geological & Environmental Sciences Cleveland State University 2121 Euclid Avenue Cleveland, OH 44115-2214, USA [email protected]

Prof. Dr. Ajit Varma Director General Amity Institute of Microbial Technology Amity University Uttar Pradesh & Vice Chairman Amity Science, Technology & Innovation Foundation Block A, Amity Campus, Sector 125 Noida, UP 201303 India [email protected]

ISSN 1613-3382 ISBN 978-3-642-14224-6 e-ISBN 978-3-642-14225-3 DOI 10.1007/978-3-642-14225-3 Springer Heidelberg Dordrecht London New York Library of Congress Control Number: 2010935854 # Springer-Verlag Berlin Heidelberg 2011 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Cover design: SPi Publisher Services Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

Current food and agriculture production appears to be sufficient to meet the present and near future demand of the world. However, due to population growth and environmental alterations that are intimately connected to global warming trends, continuous food production to meet the future demand remained uncertain. To meet additional food demand, it is crucial that we must focus on the healthy state of earth and its well being. This brings us to our relationship with the soil to which we are intimately bonded. Organic compounds, microorganisms, enzymes, and soil constitute a healthy composition for soil microecosystems. Decomposition of organic matter is central to recycling and balancing of the soil nutrients. Soil enzymes and a number of associated factors as key mediators are vital to soil macrosystem. Soil enzymes are involved in the aspects of nutrient recycling, maintaining the environmental quality, thus maintaining the soil fertility, and enhancing the productivity of economically important crops. Based on our teaching and research experience, it was imperative that a book is needed, which encompasses various facets of soil enzymes and their function in maintaining the soil quality, carbon sequestration, nutrient recycling, and variety of bioremediation of contaminated soil. In this book, we have attempted to compile broader aspects of soil enzymes including their biochemical and microbiological properties, environmental nutrients, microorganisms’ and enzymes’ mechanistic roles in maintaining a healthy soil state. The authors have presented various existing and potential environmental challenges to soil enzymes and the soil and have provided knowledge to deal with it. Numerous soil enzymes are very important to carry out basic catalytic activities and abundant biological processes in various types of soils. These processes in turn are connected to soil’s well being. The book is composed of 20 chapters encompassing various aspects of soil enzymology. The first chapter provides an overview of soil enzymes and general mechanistic aspects. Chapter 2 provides a broad and comprehensive account of the role of soil enzymes in maintaining soil health. Soil carbon sequestration and nutrient cycling is controlled by the decomposition of organic matter in the soil. Chapter 3 covers the aspects of soil enzymes, which

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Preface

facilitate soil carbon and nutrient balance. Chapter 4 highlights the enzymes that are found in forest soils, their activities, and factors affecting the activities of soil enzymes. Chapter 5 describes phosphohydrolases and their importance in organic phosphorus cycling. Hydrolytic enzymes and their role in the fast and prudent recovery of microbial cells and reestablishment of microbial communities in drywet cyclic has been presented elegantly in Chap. 6. Due to heavy machinery usage, pollution, and constant insults to environment by human interactions, the soil quality is affected in many cases irreversibly. Soil enzymes are one indicator, which corresponds to the quality and health of the soil. Chapter 7 illustrates the distinct types of soil modifications and provides relevant sources, classification, and properties of soil enzymes that make them excellent indicators. Rhizosphere is directly affected by various root secretions and its association with the soil microbes. In Chap. 8, authors present a comprehensive account of enzymes activities and factors affecting them in Rhizosphere of plant. Lignocellulose-degrading soil enzymes, phenol oxidases, and fungal oxidoreductases have been covered in Chaps. 9 through 11. Evolutionary economic principles, which modulate the production of soil enzymes, are elegantly portrayed in Chap. 12. Activity of enzymes is directly affected by temperature as determined by numerous investigations in purified laboratory systems. However, relatively fewer studies have touched upon the effect of temperature on activities of soil enzymes in its native environment. Chapter 13 fulfils this promise by providing a detailed aspect of enzyme structure–function and the effect of temperature in enzyme kinetics under field conditions. Soil enzymes, which are derived from bacterial sources including Keratinases, Pectinases, Xylanases, and Lipases, and their properties are illustrated in Chap. 14. Chapter 15 outlines the broad range of enzyme– organo-mineral interactions that occurs in the soil. Mechanistic aspects, which influence soil enzyme activity, have also been covered in the Chapter. Throughout the human history, man continued to discover methods to protect his crop from pests. Development and use of a range of pesticide has prevented total decimation of food producing crop; however, it also has caused irreversible damage to soil. While in soil, these pesticides influence the activity of soil enzymes. Chapter 16 outlines an historical and scientific perspective of the interaction of diverse pesticides with the soil and their influence on soil enzyme activity. Chapter 17 provides a unique perspective on the behavior of soil enzyme activity on volcanic soils. The chapter covers the detailed properties of volcanic ashes-derived soils and its resident soil enzymes’ activity. Screening, characterization, and optimization of microbial Pectianase have been covered in the Chap. 18. Appropriate molecular approaches in order to study polymorphism in closely related microorganisms with respect to protein phosphatase are covered in Chap. 19. In addition to use of fossil fuel and its derivative, various other pollutants including chlorinated compounds, synthetic dyes, and aromatic hydrocarbons have contaminated large areas of productive crop land. Bioremediation appears to be a logical and environmentally sustainable method to counter the effect of contaminated soil. Chapter 20 provides an in depth coverage of the production and the use of a number of white rot fungi for decontamination of oil polluted soil.

Preface

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In this volume, we attempted to cover many aspects of soil enzymology. We hope that this volume would be an essential resource for teachers, students, and research professional who are interested in basic and applied aspects of soil enzymology. In addition to extraordinary contribution by the authors of the series, the volume would have been a dream without the help of a large number of volunteers for their selfless efforts. We thank Dr. Jutta Lindenborn, Springer, Heidelberg, Germany, for her admirable patience and valuable suggestions. GCS is thankful to Dr. Bakshi for her important contribution and providing an outstanding support in fine-tuned editing of the book. Finally, we like to thank our contributors who dedicated their valuable time and expertise, and without their contribution, the volume would still be a distant dream. Cleveland, OH, USA Noida, Uttar Pradesh, India

Girish Shukla Ajit Varma

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Contents

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Soil Enzyme: The State-of-Art . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Madhunita Bakshi and Ajit Varma

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Role of Enzymes in Maintaining Soil Health . . . . . . . . . . . . . . . . . . . . . . . . . . 25 Shonkor Kumar Das and Ajit Varma

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Agricultural and Ecological Significance of Soil Enzymes: Soil Carbon Sequestration and Nutrient Cycling . . . . . . . . . . . . . . . . . . . . . 43 Wei Shi

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Enzymes in Forest Soils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 Petr Baldrian and Martina Sˇtursova´

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Extracellular Enzymes in Sensing Environmental Nutrients and Ecosystem Changes: Ligand Mediation in Organic Phosphorus Cycling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Thanh H. Dao

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Importance of Extracellular Enzymes for Biogeochemical Processes in Temporary River Sediments during Fluctuating Dry–Wet Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Annamaria Zoppini and Ju¨rgen Marxsen

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Soil Enzymes as Indication of Soil Quality . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Ayten Karaca, Sema Camci Cetin, Oguz Can Turgay, and Ridvan Kizilkaya

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Enzyme Activities in the Rhizosphere of Plants . . . . . . . . . . . . . . . . . . . . . . 149 Dilfuza Egamberdieva, Giancarlo Renella, Stephan Wirth, and Rafiq Islam

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Lignocellulose-Degrading Enzymes in Soils . . . . . . . . . . . . . . . . . . . . . . . . . . 167 Petr Baldrian and Jaroslav Sˇnajdr

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Heterophase Synthesis of Humic Acids in Soils by Immobilized Phenol Oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 A.G. Zavarzina

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Fungal Oxidoreductases and Humification in Forest Soils . . . . . . . . . . 207 A.G. Zavarzina, A.A. Lisov, A.A. Zavarzin, and A.A. Leontievsky

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Evolutionary-Economic Principles as Regulators of Soil Enzyme Production and Ecosystem Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 Steven D. Allison, Michael N. Weintraub, Tracy B. Gartner, and Mark P. Waldrop

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Controls on the Temperature Sensitivity of Soil Enzymes: A Key Driver of In Situ Enzyme Activity Rates . . . . . . . . . . . . . . . . . . . . . . 245 Matthew Wallenstein, Steven Allison, Jessica Ernakovich, J. Megan Steinweg, and Robert Sinsabaugh

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Actinomycetes: Sources for Soil Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259 V. Suneetha and Zaved Ahmed Khan

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Organo-Mineral–Enzyme Interaction and Soil Enzyme Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Andrew R. Zimmerman and Mi-Youn Ahn

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The Influence of Pesticides on Soil Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . 293 Liliana Gianfreda and Maria A. Rao

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Behavior of Enzymatic Activity in Chilean Volcanic Soil and Their Interactions with Clay Fraction . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 Analı´ Rosas, Ada Lo´pez, and Roxana Lo´pez

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Screening, Characterisation and Optimization of Microbial Pectinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 V. Suneetha and Zaved Ahmed Khan

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Molecular Techniques to Study Polymorphism between Closely Related Microorganisms in Relation to Specific Protein Phosphatase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 Rajani Malla, Utprekshya Pokharel, Ram Prasad, and Ajit Varma

Contents

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Production of Ligninolytic Enzymes by White-rot Fungi during Bioremediation of Oil-contaminated Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Natalia N. Pozdnyakova, Ekaterina V. Dubrovskaya, Oleg E. Makarov, Valentina E. Nikitina, and Olga V. Turkovskaya

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379

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Contributors

Ahn, Mi-Youn Soil and Water Science Department, University of Florida, PO Box 110290, Gainesville, FL 32611, USA, [email protected] Allison, Steven D. University of California, Irvine, 321 Steinhaus, Irvine, CA 92697, USA, [email protected] Bakshi, Madhunita Amity Institute of Microbial Technology, Amity University, Sector 125, Noida, Uttar Pradesh 201303, India, [email protected] Baldrian, Petr Laboratory of Environmental Microbiology, Institute of Microbiology of the ASCR, v.v.i., Vı´denˇska´ 1083, 14220 Praha 4, Czech Republic, [email protected] Cetin, Sema Camci Agricultural Faculty, Soil Science Department, Gaziosmanpasa University, Tokat, Turkey, [email protected] Dao, Thanh H. US Department of agriculture, Agriculture Research Service, Beltsville, MD, USA, [email protected] Das, Shonkor Kumar Department of Applied Chemistry and Biotechnology, Graduate School of Engineering, University of Fukui, 3-9-1Bunkyo, Fukui 9108507, Japan, [email protected] Dubrovskaya, Ekaterina V. Institute of Biochemistry and Physiology of Plants and Microorganisms, Russian Academy of Sciences, 13 Prospect Entuziastov, Saratov 410049, Russia, [email protected] Egamberdieva, Dilfuza Faculty of Biology and Soil Sciences, The National University of Uzbekistan, Tashkent, 100175, Uzbekistan, dilfuza_egamberdiyeva @yahoo.com

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Contributors

Ernakovich, Jessica Natural Resource Ecology Laboratory, Colorado State University, Fort Collins, CO 80523, USA; Graduate Degree Program in Ecology, Colorado State University, Fort Collins, CO 80523, USA, jagilman@nrel. colostate.edu; Gartner, Tracy B. Department of Biology and the Environmental Science Program, Carthage College, Kenosha, WI 53140, USA, [email protected] Gianfreda, Liliana Dipartimento di Scienze del Suolo della Pianta, dell’Ambiente e delle Produzioni Animali, Universita` di Napoli Federico II, Via Universita` 100, 80055 Portici, Napoli, Italy, [email protected] Islam, Rafiq Soil and Water Resources, Ohio State University South Centers, Piketon, OH, USA, [email protected] Karaca, Ayten Faculty of Agriculture, Department of Soil Science, Ankara University, Diskapi, Ankara, Turkey, [email protected] Khan, Zaved Ahmed School of Biotechnology, Chemical and Biomedical engineering, VIT University, Vellore, Tamil Nadu 632014, India, [email protected], [email protected] Kizilkaya, Ridvan Agricultural Faculty, Soil Science Department, Ondokuz Mayis University, Samsun, Turkey, [email protected] Leontievsky, A.A. Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino, Moscow Region 142292, Russia, [email protected] Lisov, A.A. Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino, Moscow Region, 142292, Russia, alex-lisov@ rambler.ru Lo´pez, Ada Instituto de Biologı´a Vegetal y Biotecnologı´a, Universidad de Talca, Curico, Chile, [email protected] Lo´pez, Roxana Facultad de Ciencias Agrarias, Departamento de Produccio´n Agrı´cola, Universidad de Talca, Curico, Chile, [email protected] Makarov, Oleg E. Institute of Biochemistry and Physiology of Plants and Microorganisms, Russian Academy of Sciences, 13 Prospect Entuziastov, Saratov 410049, Russia, [email protected] Malla, Rajani Central Department of Biotechnology, Tribhuvan University, Kathmandu, Nepal, [email protected]

Contributors

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Marxsen, Ju¨rgen Limnologische Fluss-Station des Max-Planck-Instituts fu¨r Limnologie, 36110 Schlitz, Germany; Institut fu¨r Allgemeine und Spezielle Zoologie, Tiero¨kologie, Justus-Liebig-Universita¨t Gießen, Heinrich-Buff-Ring 26-32, 35392 Gießen, Germany, [email protected] Nikitina, Valentina E. Institute of Biochemistry and Physiology of Plants and Microorganisms, Russian Academy of Sciences, 13 Prospect Entuziastov, Saratov 410049, Russia, [email protected] Pokharel, Utprekshya Department of Microbiology, Punjab University, Chandigarh, India, [email protected] Pozdnyakova, Natalia N. Institute of Biochemistry and Physiology of Plants and Microorganisms Russian Academy of Sciences, 13 Prospekt Entuziastov, Saratov 410049, Russia, [email protected] Prasad, Ram Amity Institute of Microbial Technology, Amity University Uttar Pradesh, Sector-125, New Super Express Highway, Uttar Pradesh, Noida, India, [email protected], [email protected] Rao, Maria A. Dipartimento di Scienze del Suolo della Pianta dell’Ambiente e delle Produzioni Animali, Universita` di Napoli Federico II, Via Universita` 100, 80055 Portici, Napoli, Italy, [email protected] Renella, Giancarlo Plant Nutrition and Soil Science, University of Florence, Florence, Italy, [email protected] Rosas, Anali Universidad de Concepcio´n, Facultad de Agronomı´a, Departmento de Ciencia del Suelo y Recursos Naturales, P.O. Box 537, Av. Vicente Me´ndez 595, Chillan, Chile, [email protected] Shi, Wei Department of Soil Science, North Carolina State University, Raleigh, NC 27695-7619, USA, [email protected] Sinsabaugh, Robert Department of Biology, University of New Mexico, Aluquerque, NM 87131, USA, [email protected] Sˇnajdr, Jaroslav Laboratory of Environmental Microbiology, Institute of Microbiology ASCR, Videnska 1083, 14220 Praha 4, Czech Republic, snajdr@biomed. cas.cz Steinweg, J. Megan Natural Resource Ecology Laboratory, Colorado State University, Fort Collins, CO 80523, USA; Graduate Degree Program in Ecology, Colorado State University, Fort Collins, CO 80523, USA, steinweg@warnercnr. colostate.edu

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Sˇtursova´, Martina Laboratory of Environmental Microbiology, Institute of Microbiology ASCR, Videnska 1083, 14220 Praha 4, Czech Republic, [email protected] Suneetha, V. School of Biotechnology, Chemical and Biomedical engineering, VIT University, Vellore, Tamil Nadu 632014, India, [email protected] Turgay, Oguz Can Agricultural Faculty, Soil Science Department, Ankara University, Ankara, Turkey, [email protected] Turkovskaya, Olga V. Institute of Biochemistry and Physiology of Plants and Microorganisms, Russian Academy of Sciences, 13 Prospect Entuziastov, Saratov 410049, Russia, [email protected] Varma, Ajit Amity Institute of Microbial Technology, Amity University, Sector 125, Noida 201303, Uttar Pradesh, India, [email protected] Waldrop, Mark P. U.S. Geological Survey, 345 Middlefield Rd, MS 962, Menlo Park, CA 94025, USA, [email protected] Wallenstein, Matthew Natural Resource Ecology Laboratory, Colorado State University, Fort Collins, CO 80523, USA; Graduate Degree Program in Ecology, Colorado State University, Fort Collins, CO 80523, USA, mawallen@nrel. colostate.edu; Weintraub, Michael N. Department of Environmental Sciences, University of Toledo, Toledo, OH 43606, USA, [email protected] Wirth, Stephan Leibniz-Centre for Agricultural Landscape Research, Institute of Landscape Matter Dynamics, Mu¨ncheberg, Germany, [email protected] Zavarzin, A.A. Faculty of Biology and Soil Sciences, St-Petersburg State University, 199034 St. Petersburg, Russia, [email protected] Zavarzina, A.G. Faculty of Soil Science, Moscow State University, 119991 Moscow, Russia, [email protected] Zimmerman, Andrew R. Department of Geological Sciences, University of Florida, 241 Williamson Hall, PO Box 112120, Gainesville, FL 32611, USA, aximmer@ufl. edu Zoppini, Annamaria Istituto di Ricerca Sulle Acque, Consiglio Nazionale delle Ricerche, Area della Ricerca di Roma 1-Montelibretti, via Salaria, CP10, 00016 Monterotondo, Rome, Italy, [email protected]

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Chapter 1

Soil Enzyme: The State-of-Art Madhunita Bakshi and Ajit Varma

1.1

Introduction

Soil may be defined as a thin layer of earth’s crust which serves as a natural medium for growth of plants. It is the unconsolidated mineral matter that has been subjected to and influenced by, genetic and environmental factors-parent material, climate, organisms, and topography all acting over a period of time. Soil is a natural body consisting of layers (soil horizons) of mineral constituents of variable thicknesses, which differ from the parent materials in their morphological, physical, chemical, and mineralogical characteristics. It is composed of particles of broken rock that have been altered by chemical and environmental processes that include weathering and erosion. Soil differs from its parent rock due to interactions between the lithosphere, hydrosphere, atmosphere, and the biosphere. It is a mixture of mineral and organic constituents that are in solid, gaseous, and aqueous states. Soils differ among themselves in some or all the properties, depending on the differences in the genetic and environmental factors. Thus some soils are red, some are black, some are deep and some are shallow, some are coarse textured and some are fine-textured. They serve as a reservoir of nutrients and water for crops, provide mechanical anchorage and favorable tilth. The components of soil are mineral matter, organic matter, water, and air, the proportions of which vary and which together form a system for plant growth; hence the need to study the soils in perspective.

M. Bakshi and A. Varma (*) Amity Institute of Microbial Technology, Amity University Uttar Pradesh, Sector 125, Noida 201303, India e-mail: [email protected]

G. Shukla and A. Varma (eds.), Soil Enzymology, Soil Biology 22, DOI 10.1007/978-3-642-14225-3_1, # Springer-Verlag Berlin Heidelberg 2011

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1.2

Soil: A Physical Point of View

The physical properties of a soil like the water holding capacity, permeability to water, aeration, plasticity, and nutrient-supplying ability, are influenced by the size, proportion, arrangement, and mineral composition of the soil particles.

1.2.1

Particle Shape and Size

The soil particles vary in shape from spherical to angular. They differ in size from gravel and sand to fine clay (Table 1.1)

1.2.2

Textural Classes

The varying proportion of particles of different size groups in a soil constitute what is termed as a textural class. The principal textural classes are: clay, sandy clay, silty clay, clay loam, sandy clay loam, silty clay loam, loam, sandy loam, silt loam, sand, loamy sand, and silt. The textural classes differ not only in the particle size analysis, but also in their bearing on some of the important factors affecting plant growth, such as 1. 2. 3. 4.

The moveability and availability of water Aeration Workability The contents of plant nutrients

Sandy soils are very permeable and well drained but are less water retentive and hence need more frequent irrigation for successful crop growth than fine textured soils. The clayey soils can hold more moisture, but they have high wilting percentage. The rate of water intake of these soils is low. They are subject to water-logging resulting in poor aeration and workability. The moderately fine-textured soils for Table 1.1 Classification of Soil according to their particle size

Types Gravel Very coarse sand Coarse sand Medium sand Fine sand Very fine sand Silt Clay

International system of classification (Diameter in mm) 2 and more 2–1 1.0–0.5 0.5–0.25 0.25–1 0.10–0.05 0.05–0.02 less than 0.002

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example, loams, clay loams or silt loams are by far the excellent soils for plant growth, since they have the advantages of both sands and clays.

1.2.3

Colors

Colors of a soil are indicator of soil conditions and some important properties. It is due either to mineral or organic matter or mostly to both. Red, yellow, or brown colors are usually related to the different degrees of oxidation, hydration, and diffusion of iron oxides in the soil. Dark colors of a soil are associated with one or a combination of several factors, including impeded drainage conditions, content, and state of decomposition of organic matter.

1.2.4

Density

Soils having larger particles are usually heavier in weight per unit volume than those having smaller particles. True density of a soil is based on the individual densities of soil constituents and according to their proportionate contribution.

1.2.5

Pore Space

The pore space of soil is the portion occupied by air and water and it is determined largely by structural conditions. Sands have low pore space of about 30%, whereas clays may have as much as 50–60%. Although clays possess greater total porosity than the sands the pore spaces in the latter being individually larger are more conducive to good drainage and aeration.

1.2.6

Plasticity and Cohesion

Plasticity and cohesion reflect the soil consistency and workability of the soils. Plasticity is the property that enables a moist soil to change shape on the application of force and retain this shape even when the force is withdrawn. Cohesion is the tendency of the particles to stick to one another. Plastic soils are cohesive. On this basis, sandy soils may be considered to be non-plastic and clayey soils to be plastic.

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1.2.7

Soil Temperature and Heat

Soil temperature is one of the important factors that control the microbiological activity and all the processes involved in the growth of plants. The temperature needed for germination and root growth varies with crops and varieties. Crops, for example, wheat, barley, and peas grown in India during winter germinate at relatively low temperatures as compared with maize, and those at which groundnut or cotton germinate. Heat is necessary for seed germination, root growth, and other biological activities.

1.2.8

Soil Water

Water has perhaps the greatest influence on the growth and yield of a crop. It is needed in much larger quantity than that of any other substance that contributes to growth or yield. Water serves the following functions in relation to plant life. 1. It is an essential part of plant food. It constitutes nearly 90% of plant tissues. 2. It serves as a solvent and carrier of plant nutrients. 3. It maintains cell turgidity and regulates temperature. Water is held in soil in the following forms: 1. Hygroscopic Water: It occurs as a thin film (4–5 mm) and is held tenaciously with a tension of 31 atmospheres or more. It is not available to plants. 2. Capillary Water: It forms a continuous film around soil particles (outside the film of hygroscopic water) and in the micropore spaces. It is held by surface tension. Capillary water is held at a tension ranging from 1/3 to 31 atmospheres. 3. Gravitational Water: It is free water held at a tension below 1/3 atmospheres. It saturates the soil and percolates downwards under the influence of gravity.

1.3 1.3.1

Soil: A Chemical Point of View Organic Matter

Organic matter, though forming a small part of mineral soils, plays a vital role in the productivity and conditioning of soils. It serves as a source of food for soil bacteria and fungi, which are responsible for converting complex organic materials into simple substances readily used by the plants. The intermediate products of decomposition of fresh organic matter help to increase the physical condition of the soil. The addition of organic matter also improves the working quality or friability of the soil. In association with clay and calcium, it helps to form the aggregates of soil particles to produce the “crumb structure”.

1 Soil Enzyme: The State-of-Art

1.3.2

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Humus

The organic matter in the soil consists largely of plant remains, the residues of soilmicroorganisms feeding on them, and several products of their decomposition. In consonance with its origin, organic matter contains most of the mineral elements found in the plants. The plant remains may occur in recognizable form but more commonly they are found as a fairly stable, dark, amorphous complex colloidal substance called humus. Chemically, humus represents a mixture of decomposed or altered products of carbohydrates, proteins, fats, resins, wax, and other similar substances. These complex compounds are gradually decomposed by soil organisms into simple mineral salts, carbon dioxide, water, organic acids, ammonia, methane, and free nitrogen, depending upon the initial composition of the organic matter. The average composition of humus is as follows Table 1.2.

1.3.3

Soil Colloids

The most active portions of the soil are those which are in the colloidal state. The colloidal state implies a two-phase system in which the material called the dispersed phase (fine clay and humus) is dispersed in the dispersed medium (water). In soils, the mineral and organic colloids exist in intimate and heterogeneous admixture. The mineral colloid is present almost exclusively as the clay of various kinds, whereas the organic colloid is present as humus. It is generally believed that clay particles less than one micron in diameter possess colloidal properties and these properties increase with a decrease in the size of the particles. The most distinctive colloidal properties are: 1. The large specific surface or interface and 2. The capacity to hold solids, gases, salts, and ions.

1.4

Soil: A Biological Point of View

Soil is a natural medium in which microbes live, multiply, and die. Microbial diversity in the soil is a critical environmental topic that concerns people from all Table 1.2 Chemical composition of humus

Carbon Oxygen Nitrogen Hydrogen Ash (containing phosphorus, potassium, sulfur and other elements)

Percentage 50 35 5 5 5

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walks of life. Raw organic matter in the soil is not directly used by the plants as food. It must be broken down first into humus and then into simpler products before it can be utilized. This work is done by different kinds of microorganisms in the soil. The decomposition of organic matter forms part of the feeding and growth process of these microscopic plants and animals. However, not all the soil organisms are beneficial. There are certain bacteria which under anaerobic conditions of water-logged soil cause denitrification releasing free nitrogen which gets lost into the air.

1.4.1

Kind of Soil Organisms

Soil contains a wide range of microorganisms described as “black box” (Paul and Clark 1989). Microorganisms in soil are closely associated with soil particles, mainly clay-organic matter complexes (Foster 1988). Their activity and interaction with other microbes and larger organisms and with soil particles depend largely on conditions at the microhabitat level that may differ among microhabitats even over very small distances (Wieland et al. 2001). Increasing attention is being drawn to microorganisms because the fertility of soil depends not only on its chemical composition, but also on the qualitative and quantitative nature of microorganisms inhabiting it (Fig. 1.1; Giri et al. 2005). Microorganisms are generally divided into five major taxonomic categories: Algae, Eubacteria, Fungi, Protists, and Viruses. The more complex cells constitute eukaryotes, which include Algae, Fungi, and Protists. The less complex cell constitutes prokaryotes comprising two microbial groups: Eubacteria (including Cyanobacteria) and the Archaebacteria.

1.4.1.1

Eubacteria

Eubacteria also known as true bacteria are recognized as the most dominant group of microorganisms among the various kinds of soil (Liesack and Stackebrandt 1992; Visscher et al. 1992; Borneman et al. 1996).They are present in all types of soil, but their population decreases as the depth of soil increases. Bacteria live in soil as cocci (sphere, 0.5 mm), bacilli (rod, 0.5–0.3 mm), or spiral. The bacilli are common in soil, whereas spirilli are very rare in natural environments (Giri et al 2004). The most common bacteria belong to the genera Pseudomonas, Arthrobacter, Clostridium, Achromobacte, Bacillus, Micrococcus, Flavobacterium, Coryne bacterium, Sarcina, Azospirillum, and Mycobacteria (Loper et al. 1985; Bruck 1987; Lynch 1987a, b). Escherichia is encountered rarely in soil except as a contaminant from sewage, whereas Aerobacter is frequently encountered and is probably a normal inhabitant of certain soils (Subba Rao 1997). Cyanobacteria are organisms that are very important in the formation of biological soil crusts (Fig.1.2). They are photosynthetic and live within the first

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Fig. 1.1 A typical view of mycorrhizosphere (a) an enlarged view of the root system containing the dynamic root system embedded with nanoparticles (b)

Fig. 1.2 Cyanobacteria present on soil surface

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ten inches of topsoil. Cyanobacteria help to reduce erosion by helping bind the particles of soil together. When the filaments of cyanobacteria become wet, they absorb water and swell up to ten times their original size which helps to store moisture within the upper layer of soil where many plants root systems and other organisms live. Cyanobacteria also play a more direct role in aiding plant survival and growth. Cyanobacteria live in the roots of some plants viz., Bryophytes (species of Anthoceros, Riccia, and Marchantia), Pteridophytes (Azolla spp) and coralloid roots in Cycas spp. (Gymnosperms). These symbioses provide dinitrogen directly for the hosts use. They have also formed symbiotic relationships with fungus (Schussler and Kluge 2001) and lichens (Kranner et al. 2002).

1.4.1.2

Fungi

Soil fungi are microscopic plant-like cells that grow in long threadlike structures or hyphae that make a mass called mycelium. The mycelium absorbs nutrients from the roots it has colonized, surface organic matter or the soil. Fungi perform important services related to water dynamics, nutrient cycling, and disease suppression. There are three functional groups of fungi found in soil:

Decomposers Decomposers or saprophytic fungi convert dead organic matter into fungal biomass (i.e., their own bodies), carbon dioxide, and organic acids. They are essential for the decomposition of hard woody organic matter. By consuming the nutrients in the organic matter, they play an important role in immobilizing and retaining nutrients in the soil. The organic acids they produce as by products help create organic matter that is resistant to degradation.

Mutualists These fungi develop mutually beneficial relationships with plants, for example, Mycorrhiza – P. indica, which colonize plant roots where they help the plant to obtain nutrients such as phosphorus from the soil. Their mass hides the roots from pests and pathogens and provides a greater root area, through which the plant can obtain nutrients.

Pathogens This group includes the well known fungi such as Verticillium, Phytophthora, Rhizoctonia, and Pythium. These organisms penetrate the plant and decompose

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the living tissue, creating a weakened nutrient deficient plant or death. The pathogenic fungi are usually the dominant organism in the soil.

1.4.1.3

Algae

Algae are generally found on the surface of moist soils, where there is sufficient light for their photosynthetic reactions. The major types present are green algae Chlorophyceae, and the diatoms (Bacillariophyceae). Microalgae are tiny plants which are active converters of solar energy, carbon dioxide, and other nutrients into sugars, proteins, and other complex organic compounds beneficial to the nutrient cycling and soil structure of croplands. Some species are able to fix atmospheric nitrogen; others form compounds conducive to the growth of associated plants. Some blue-green algae, along with a few other species (such as those of the genus Azotobacter), are rather unique in their ability to perform the nitrogen-fixing function in the presence of oxygen at the soil/ air interface, and excrete substances which further serve to glue soil particles into aggregates. This provides more aerated soil area for further growth of – these and associated beneficial microorganisms. Additionally, Anabaena has been shown to help release soil-bound phosphorus, although the exact ways in which this happens are not known at this time. Some of the common green algae occurring in most soils belong to the genera Chlorella, Chlamydomonas, Chlrococcum, Oedogonium, Chlorochytrium, and Protosiphone.

1.4.2

Population Density

The soil organisms vary in number from a few per hectare to many millions per gram of soil. The density of population is determined by food supply, moisture, temperature, physical condition, and the reaction of the soil. In neutral soils, bacteria dominate over other types of microscopic life. If the soil is acidic and rich in organic matter, fungi predominate. Algae abound on the soil in constantly moist or shady situations.

1.4.3

Bacterial Activity

The soil microflora typically produces ammonia from organic compounds when they set free more of nitrogen than they can assimilate and convert into their own protoplasm. Ammonia so released is converted into nitrites by one group of organisms called Nitrosomonas: NH4þ þ 11=2O2 ! NO2   þH2 O þ 2Hþ

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The nitrite can then be further oxidized by Nitrobacter and Nitrococcus to yield nitrate. NO2   þ1=2O2 ! NO3 When two genera work together, ammonia in the soil is oxidized to nitrate in a process called nitrification (Prescott’s Microbiology, 3rd ed, 1996).

1.4.3.1

Energetics of Nitrification

Energy released upon the oxidation of both ammonia and nitrite is used to make ATP by oxidative phosphorylation. Since electron moves spontaneously from molecules of more negative reduction potential to acceptors with more positive reduction potential, so molecules like ammonia and nitrite which have more positive reduction potentials than NAD+, cannot directly donate their electrons to form the required NADH and NADPH. Nitrifying bacteria solve this problem by using proton motive force or ATP to reverse the flow of electrons in their electron transport chains and reduce NAD+ with electrons from nitrogen. Since energy is used to generate NADH as well as ATP, the net yield is fairly low (Fig. 1.3). The nitrate-forming bacteria are generally confined to the top 25–30 cm of the soil, where the content of organic matter is also more. These organisms are most active between 25 and 38 C and under favorable conditions of tillage, aeration, – 0.32V NADH+H+ NAD+

Flavoprotein

–

NO2

ADP + Pi ATP

Cytochromes

Cyt a1

Cyt a3

Fig. 1.3 Reverse electron flow. The flow of electrons in the transport chain of Nitrobacter (Prescott et al., 5ed)

1/2O2

H2O +0.82V

ATP+Pi ADP

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neutral soil reaction, and moisture content at field capacity. The bacteria cease or reduce their activity when the pH value of the soil falls below 5.0.

1.5

Enzymes

Enzymes are biologically produced proteinic substances, having specific activation in which they combine with their substrates in such a stereoscopic position that they cause changes in the electronic configuration around certain susceptible bonds. Their significance in all spheres including soil is worth tested and reported. In plant nutrition, their role cannot be substituted by any other substance and their function is quite pragmatic in solubilizing and dissolving the much needed food in ionic forms for the very survival of the animal and plant kingdom. Enzymes are the key to understanding below-ground biochemistry and the role of soil in the global carbon cycle. These biological mediators of change are active both within living soil organisms and independently as extracellular proteins that are actively secreted into the soil by roots and fungi, or released as prokaryotic and eukaryotic cells that die and decompose. These enzymes can persist in the soil for weeks while maintaining a ghostly after life activity.

1.6

Soil Enzymes

Soil enzymes play key biochemical functions in the overall process of organic matter decomposition in the soil system (Burns 1983; Sinsabaugh et al. 1991). They are important in catalyzing several important reactions necessary for the life processes of micro-organisms in soils and the stabilization of soil structure, the decomposition of organic wastes, organic matter formation, and nutrient cycling (Dick et al. 1994). These enzymes are constantly being synthesized, accumulated, inactivated, and/or decomposed in the soil, hence playing an important role in agriculture and particularly in nutrients cycling (Tabatabai 1994; Dick 1997). A better understanding of the role of these soil enzymes activities in the ecosystem will potentially provide a unique opportunity for an integrated biological assessment of soils due to their crucial role in several soil biological activities, their ease of measurement, and their rapid response to changes in soil management practices (Dick 1994, 1997; Bandick and Dick 1999). The activity of these enzymes in soil undergoes complex biochemical processes consisting of integrated and ecologically-connected synthetic processes, and in the immobilization and enzyme stability (Khaziyev and Gulke 1991). In this regard, all soils contain a group of enzymes that determine soil metabolic processes (McLaren 1975) which, in turn, depend on its physical, chemical, microbiological, and biochemical properties.

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Soil enzyme activities are often used as indices of microbial growth and activity in soils. Quantitative information concerning which soil enzymes most accurately reflect microbial growth and activity is lacking (Department of Soil and Environmental Sciences, University of California, USA). Categories of soil enzymes include: 1. Enzymes associated with living, metabolically active cells in soil; found in cell’s cytoplasm, bound to cell wall or as extracellular enzymes that have been recently produced by the cell (Fig. 1.4). 2. Enzymes associated with viable but non-proliferating cells (such as spores) 3. Enzymes that are attached to dead cells or to cell debris, or which have diffused away from dead/dying cells that originally produced them. 4. Enzymes that are “permanently” immobilized on soil clay and humic colloids. Such enzymes can remain active for long periods of time. Such immobilized soil enzymes can arise from either eukaryotic or prokaryotic cells. Binding of enzymes to soil surfaces (especially clay and humic materials) may take place via Ionic interactions, Covalent bonds, Hydrogen bonding, Entrapment of enzyme by soil colloids, and other mechanisms (Fig. 1.5). These enzymes may include amylase, arylsulphatases, glucosidase, cellulose, chitinase, dehydrogenase, phosphatase, protease, and urease released from plants (Miwa et al. 1937), animals (Kanfer et al. 1974), organic compounds and microorganisms (Dick and Tabatabai 1984; James et al. 1991; Richmond 1991; Hans and Snivasan 1969; Shawale and Sadana 1981), and soils (Cooper 1972; Gupta et al. 1993; Ganeshamurthy et al. 1995).

organic substrates

enzyme

clay particles

Fig. 1.4 A soil extra cellular enzyme physically protected by a clay particle sheath

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Fig. 1.5 Enzymes associated with clay minerals via different types of bond

1.6.1

Amylase

Amylase is a starch hydrolyzing enzyme (Ross 1976). It is known to be constituted by amylase and amylase (Pazur 1965; King 1967; Thoma et al. 1971). Studies have shown that amylases are synthesized by plants, animals, and micro-organisms, whereas amylase is mainly synthesized by plants (Pazur 1965; Thoma et al. 1971). This enzyme is widely distributed in plants and soils so it plays a significant role in the breakdown of starch. The roles and activities of amylase and amylase enzymes may be influenced by different factors ranging from cultural practices, type of vegetation, environment, and soil types (Ross 1968; Ross and Roberts 1970; Pancholy and Rice 1973; Ross 1975). For example, plants may influence the amylase enzyme activities of soil by directly supplying enzymes from their residues or excreted compounds, or indirectly providing substrates for the synthetic activities of micro-organisms.

1.6.2

b-Glucosidase

b-Glucosidase is a common and predominant enzyme in soils (Eivazi and Tabatabai 1988; Tabatabai 1994). It is named according to the type of bond that it hydrolyses. This enzyme plays an important role in soils because it is involved in catalyzing the hydrolysis and biodegradation of various glucosides present in plant debris decomposing in the ecosystem (Ajwa and Tabatabai 1994; Martinez and Tabatabai 1997). Its final product is glucose, an important C energy source of life to microbes in the soil (Esen 1993). b-Glucosidase is characteristically useful as a soil quality indicator, and may give a reflection of past biological activity, the capacity of soil to stabilize the soil organic

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matter, and can be used to detect management effect on soils (Bandick and Dick 1999; Ndiaye et al. 2000). b-Glucosidase enzyme is very sensitive to changes in pH, and soil management practices (Dick et al. 1996; Acosta-Martı´nez and Tabatabai 2000; Kuperman and Carreiro 1997; Bergstrom et al. 1998; Leiro´s et al. 1999; Bandick and Dick 1999; Madejo´n et al. 2001). Acosta-Martı´nez and Tabatabai (2000) reported b-glucosidase as sensitive to pH changes. b-Glucosidase enzyme is also known to be inhibited by heavy metal contamination such as Cu and several others (Haanstra and Doelman 1991; Deng and Tabatabai 1995; Wenzel et al. 1995).

1.6.3

Cellulase

Cellulose is the most abundant organic compound in the biosphere, comprising almost 50% of the biomass synthesized by photosynthetic fixation of CO2 (Eriksson et al. 1990). The growth and survival of micro-organisms that are important in most agricultural soils depend on the carbon source contained in the cellulose occurring in the soils (Deng and Tabatabai 1994). Cellulose in plant debris has to be degraded into glucose, cellobiose and high molecular weight oligosaccharides by cellulase enzymes (White 1982). Cellulases are a group of enzymes that catalyze the degradation of cellulose; polysaccharides build up of four linked glucose units (Deng and Tabatabai 1994). Currently, it is accepted that the cellulase system comprises three major types of enzymes. They are: endo-1,4-b-glucanase which attacks the cellulose chains at random, exo-1,4-bglucanase which removes glucose or cellobiose from the non-reducing end of the cellulose chains, and b-D-glucosidase which hydrolyses cellobiose and other water soluble cellodextrins to glucose. Studies have shown that activities of cellulases in agricultural soils are affected by several factors. These include temperature, soil pH, water, and oxygen contents (abiotic conditions), the chemical structure of organic matter and its location in the soil profile horizon (Rubidge 1977; Gomah 1980; Tabatabai 1982; Klein 1989; Deng and Tabatabai 1994; Alf and Nannipieri 1995), quality of organic matter/plant debris and soil mineral elements (Burns 1978; Hope and Burns 1987; Klein 1989; Sinsabaugh and Linkins 1989; Deng and Tabatabai 1994) and the trace elements from fungicides (Deng and Tabatabai 1994; Petker and Rai 1992; Arinze and Yubedee 2000; Atlas et al. 1978; Vincent and Sisler 1968). Since cellulases play an important role in global recycling of the most abundant polymer, cellulose in nature, it would be of critical importance to understand this enzyme better so that it may be used more regularly as a predictive tool in our soil fertility programmes.

1.6.4

Laccase

Laccases (EC. 1.10.3.2 p-benzenediol: oxygen oxidoreductase) belong to a family of multi-copper oxidases. Laccases are widely distributed enzymes in higher plants,

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fungi, some insects, and bacteria. They are characterized by low substrate specificity, oxidizing various substrates, including diphenols, polyphenols, different substituted phenols, diamines, aromatic amines, and even inorganic compounds like iodine. Laccases oxidize their substrates by a one-electron oxidation mechanism, and they use molecular oxygen as an electron acceptor. The enzyme having pH optimum within 5–8 and working at temperatures between 30 and 80 C, is well suited to industrial applications requiring high pH and temperature conditions whereas the majority of known fungal laccases function in an acidic pH range and are not very thermostable. The exo-oxidative enzyme laccase has been detected in a large number of basidiomycete ectomycorrhizal fungi, in a few ectomycorrhizal ascomycetes and only in one endomycorrhizal species. Most fungi tested showed a strong laccase activity. Piriformospora indica and Sebacina vermifera ss. Warcup and Talbot also showed a positive reaction to the ABTS test i.e., presence of laccase activity (Pham et al. 2004).

1.6.5

Chitinase

Chitinase or chitinolytic enzymes are key enzymes responsible for the degradation and hydrolysis of chitin (poly b-1-4-(2-ncetamido-2-deoxy)-D-glucoside). They are also considered as the major structural component of many fungal cell walls that use the hyper parasitism mechanisms against pests/pathogen attack, (BartinickiGarcia 1968; Chet and Henis 1969; Chet and Henis 1975; Chet 1987). For example, in plants, the chitinase enzyme is induced and accumulated in response to microbial infections and it is thought to be involved in the defense of plants against pathogen infections (Boiler et al. 1983; Boiler 1985). As biological control of most pathogenic diseases is increasingly gaining popularity in recent times due to their environmental friendliness, a better understanding of the chitinolytic enzymes is likely to uncover more application avenues for this enzyme in agricultural systems and, consequently, increase plant growth and final yields.

1.6.6

Dehydrogenase

The dehydrogenase enzyme activity is commonly used as an indicator of biological activity in soils (Burns 1978).This enzyme is considered to exist as an integral part of intact cells but does not accumulate extracellularly in the soil. Dehydrogenase enzyme is known to oxidize soil organic matter by transferring protons and electrons from substrates to acceptors. These processes are part of respiration pathways of soil microorganisms and are closely related to the type of soil and soil air-water conditions (Doelman and Haanstra 1979; Kandeler 1996; Glinski and Stepniewski 1985).

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Studies on the activity of dehydrogenase enzyme in the soil are very important as it may give indications of the potential of the soil to support biochemical processes which are essential for maintaining soil fertility. A study by Brzezinska et al. (1998) suggested that soil water content and temperature influence dehydrogenase activity indirectly by affecting the soil redox status. The relationship between dehydrogenase activity and redox potential (Eh) as well as Fe2+ content may also be used to illustrate the reactions of soil microorganisms to the changes in soil environment. Additionally, dehydrogenase enzyme is often used as a measure of any disruption caused by pesticides, trace elements, or management practices to the soil (Reddy and Faza 1989; Wilke 1991; Frank and Malkomes 1993), as well as a direct measure of soil microbial activity (Skujins 1978; Trevors 1984; Garcia and Herna´ndez 1997). It can also indicate the type and significance of pollution in soils. For example, dehydrogenase enzyme is high in soils polluted with pulp and paper mill effluents (McCarthy et al. 1994) but low in soils polluted with fly ash (Pitchel and Hayes 1990).

1.6.7

Phosphatase

Phosphatases are a broad group of enzymes that are capable of catalyzing hydrolysis of esters and anhydrides of phosphoric acid (Schmidt and Laskowski 1961). Apart from being good indicators of soil fertility, phosphatase enzymes play key roles in the soil system (Dick and Tabatai 1992; Dick et al. 2000). In fungi, these phosphatases may be located in the periplasmic space, cell wall, vacuoles, and culture medium. Acid and alkaline phosphatases are the two forms of active phosphatase. Alkaline phosphatase, occurs in roots mainly after mycorrhizal colonization and has been proposed as a marker for analyzing the symbiotic efficiency of root colonization (Tisserant et al. 1993).A newly discovered fungus Piriformospora indica is known to produce one form of intracellular ACPase irrespective of the phosphate concentration. The enzyme is possibly a constitutive enzyme showing a molecular mass of 66 kDa (Malla et al. 2007). In soil ecosystems, these enzymes are believed to play critical roles in P cycles (Speir and Ross 1978) as evidence shows that they are correlated to P stress and plant growth. The amount of acid phosphatase exuded by plant roots has been shown to differ between crop species and varieties, (Ndakidemi 2006; Izaguirre-Mayoral et al. 2002) as well as crop management practices (Ndakidemi 2006; Patra et al. 1990; Staddon et al. 1998; Wright and Reddy 2001). For instance, research has shown that legumes secrete more phosphatase enzymes than cereals (Yadav and Tarafdar 2001). This may probably be due to a higher requirement of P by legumes in the symbiotic nitrogen fixation process as compared to cereals. In their studies, Li et al. (2004) reported that chickpea roots were also able to secrete greater amounts of acid phosphatase than maize.

1 Soil Enzyme: The State-of-Art

1.6.8

17

Protease

Proteases in soil play a significant role in N mineralization (Ladd and Jackson 1982), an important process regulating the amount of plant available N (Stevenson 1986) and plant growth. This enzyme in the soil is generally associated with inorganic and organic colloids (Burns 1982; Nannipieri et al. 1996). Protease activities have been reported to occur partly in soil as a humocarbohydrate complex (Mayaudon et al. 1975; Batistic et al. 1980). The amount of this extracellular enzyme activity may be indicative not only of the biological capacity of soil for the enzymatic conversion of the substrate, which is independent of the extent of microbial activity, but might also have an important role in the ecology of micro-organisms in the ecosystem (Burns 1982).

1.6.9

Urease

Urease enzyme is responsible for the hydrolysis of urea fertilizer applied to the soil into NH3 and CO2 with the concomitant rise in soil pH (Andrews et al. 1989; Byrnes and Amberger 1989). Due to this role, urease activities in soils have received a lot of attention since it was first reported by Rotini (1935), a process considered vital in the regulation of N supply to plants after urea fertilization. Urease extracted from plants or micro-organisms is rapidly degraded in soil by proteolytic enzymes (Burns et al. 1972; Pettit et al. 1976; Zantua and Bremner 1977). Urease activity in soils is influenced by many factors. These include cropping history, organic matter content of the soil, soil depth, soil amendments, heavy metals, and environmental factors such as temperatures (Tabatabai 1977; Bremner and Mulvaney 1978; Yang et al. 2006). Generally, urease activity increases with increasing temperature. It is suggested that higher temperatures increase the activity coefficient of this enzyme. Therefore, it is recommended that urea be applied at times of the day when temperatures are low. This is because during such times the activation energy is low, thus, resulting in minimum loss of N by the volatilization process. A better understanding of this enzyme would provide more effective ways of managing urea fertilizer especially in high rainfall areas, flooded soils, and irrigated lands as well as where urea fertilizer is vulnerable to urease enzyme.

1.6.10 Arylsulphatase Sulfur uptake in plants is in the form of inorganic sulfate (SO4) and its availability depends on its mineralization or mobilization (Williams 1975; Fitzgerald 1976) from aromatic sulfate esters (RO–SO3). This is due to the fact that certain proportions of sulfur in different soil profiles are bound into organic compounds and are indirectly available to plants.

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In this regard, its availability will depend on the extracellular hydrolysis of these aromatic sulphate esters or intracellular oxidation of soluble organic matter absorbed by the micro-organisms to yield energy and carbon skeletons for biosynthesis by which some SO4–S are released as a by-product (Dodgson et al. 1982).

1.7

Conclusion

Physical, chemical, and biological properties of a soil are indicators of soil quality, while soil fertility can be determined by its biological activity. Soil provides natural environment for the survival of microorganism and they need favorable physical and chemical conditions for their optimal function. An imbalance of soil microorganisms, nutrient deficiency, and change in physicochemical properties, like a decrease in pH, can result in decreased soil fertility and crop productivity. Understanding other possible roles of soil enzymes is vital to soil health and fertility management in ecosystems. These enzymes may have significant effects on soil biology, environmental management, growth, and nutrient uptake in plants growing in ecosystems.

References Acosta-Martı´nez V, Tabatabai MA (2000) Enzyme activities in a limed agricultural soil. Biol Fertil Soils 31:85–91 Ajwa HA, Tabatabai MA (1994) Decomposition of different organic materials in soils. Biol Fertil Soils 18:175–182 Alf K, Nannipieri P (1995) Cellulase activity, Methods in Applied Soil Microbiology and Biochemistry. Academic Press, London Andrews RK, Blakeley RL, Zerner B (1989) Urease: a Ni (II) metalloenzyme. In: Lancaster JR (ed) The bioinorganic chemistry of nickel. VCH Publishers, New York, pp 141–166 Arinze AE, Yubedee AG (2000) Effect of fungicides on Fusarium grain rot and enzyme production in maize (Zea mays L.). Glob J Appl Sci 6:629–634 Atlas RM, Pramer D, Bartha R (1978) Assessment of pesticide effects on non-target soil microorganisms. Soil Biol Biochem 10:231–239 Bandick AK, Dick RP (1999) Field management effects on soil enzyme activities. Soil Biol Biochem 31:1471–1479 Bartinicki-Garcia S (1968) Cell wall chemistry, morphogenesis and taxonomy of fungi. Annu Rev Microbiol 144:346–349 Batistic L, Sarkar JM, Mayaudon J (1980) Extraction, purification and properties of soil hydrolases. Soil Biol Biochem 12:59–63 Bergstrom DW, Monreal CM, King DJ (1998) Sensitivity of soil enzyme activities to conservation practices. Soil Sci Soc Am J 62:1286–1295 Boiler T (1985) Induction of hydrolases as a defense reaction against pathogens. In: Key JL, Kosuge T (eds) Cellular and molecular biology of plum stress. Liss, New York, pp 247–262 Boiler T, Gehri A, Mauch F, Vogeli U (1983) Chitinase in bean leaves: induction by ethylene, purification, properties and possible function. Planta 157:22–31

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Borneman J, Skroach PW, O’Sullivian EW, Palus JA, Rumjanek NG, Jansen JL, Nienhuis J, Triplett EW (1996) Molecular microbial diversity of an agricultural soil in Wisconsin. Appl Environ Microbiol 62:1935–1943 Bremner JM, Mulvaney RL (1978) Urease activity in soils. In: Bums RG (ed) Soil enzymes. Academic, London, pp 149–196 Bruck TD (1987) The study of microorganisms in situ: progress and problems. In: Fletcher M, Gray TRG, Jones JG (eds) Ecology of microbial communities. SGM symposium 41. Cambridge University Press, Cambridge, pp 455–493 Brzezinska M, Stepniewska Z, Stepniewski W (1998) Soil oxygen status and dehydrogenase activity. Soil Biol Biochem 30(13):1783–1790 Burns RG (1978) Enzyme activity in soil: some theoretical and practical considerations. In: Bums RG (ed) Soil enzymes. Academic, London, pp 295–340 Burns RG (1982) Enzyme activity in soil: location and possible role in microbial ecology. Soil Biol Biochem 14:423–427 Burns RG (1983) Extra cellular enzyme–substrate interactions in soil. In: Slater JH, Wittenbury R, Wimpenny JWT (eds) Microbes in their natural environment. Cambridge University Press, London, pp 249–298 Burns RG, Pukite AH, McLaren AD (1972) Concerning the location and persistence of soil urease. Soil Sci Soc Am Proc 36:308–311 Byrnes BH, Amberger A (1989) Fate of broadcast urea in a flooded soil when treated with N-(n-butyl) thiophospheric triamide, a urease inhibitor. Fertil Res 18:221–231 Chet I (1987) Trichoderma-application, mode of action, and potential as biocontrol agent of soil borne pathogenic fungi. In: Chet I (ed) Innovative approaches to plant disease control. Wiley, New York, pp 137–349 Chet I, Henis Y (1969) Effect of catechol and disodium EDTA on melanin content of hyphal and sclerotial walls of Sclerotium rolfsii Sacc. and the role of melanin in the susceptibility of these walls to _-l-3 glucanase and chitinase. Soil Biol Biochem 1:131–138 Chet I, Henis Y (1975) Sclerotial morphogenesis in fungi. Annu Rev Phytopathol 13:169–192 Cooper PJM (1972) Arylsulphatase activity in Northern Nigerian soils. Soil Biol Biochem 4:333–337 Deng SP, Tabatabai MA (1994) Cellulase activity of soils. Soil Biol Biochem 26:1347–1354 Deng SP, Tabatabai MA (1995) Cellulase activity of soils: effect of trace elements. Soil Biol Biochem 27:977–979 Dick RP (1994) Soil enzyme activities as indicators of soil quality. In: Doran JV, Coleman DC, Bezdicek DF, Stewart BA (eds) Defining soil quality for a sustainable environment, soil science society of America. American Society of Agriculture, Madison, pp 107–124 Dick RP (1997) Soil enzyme activities as integrative indicators of soil health. In: Pankhurst CE, Doube BM, Gupta VVSR (eds) Biological indicators of soil health. CAB International, Wellingford, pp 121–156 Dick WA, Tabatabai MA (1984) Kinetic parameters of phosphatase in soils and organic waste materials. Soil Sci 137:7–15 Dick WA, Tabatai MA (1992) Potential uses of soil enzymes. In: Metting FB Jr (ed) Soil microbial ecology: applications in agricultural and environmental management. Marcel Dekker, New York, pp 95–127 Dick RP, Sandor JA, Eash NS (1994) Soil enzyme activities after 1500 years of terrace agriculture in the Colca Valley, Peru. Agric Ecosyst Environ 50:123–131 Dick RP, Breakwell DP, Turco RF (1996) Soil enzyme activities and biodiversity measurements as integrative microbiological indicators. In: Methods for assessing soil quality. Soil Science Society of America. Madison, WI, pp 9–17 Dick WA, Cheng L, Wang P (2000) Soil acid and alkaline phosphatase activity as pH adjustment indicators. Soil Biol Biochem 32:1915–1919 Dodgson KS, White G, Fitzgerald JW (1982) Sulphatase enzyme of microbial origin, vol I. CRC, Boca Raton, FL

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Doelman P, Haanstra L (1979) Effect of lead on soil respiration and dehydrogenase activity. Soil Biol Biochem 11:475–479 Eivazi F, Tabatabai MA (1988) Glucosidases and galactosidases in soils. Soil Biol Biochem 20:601–606 Eriksson KEL, Blancbette RA, Ander P (1990) Biodegration of cellulose. In: Eriksson KEL, Blanchette RA, Ander P (eds) Microbial and enzymatic degradation of wood and wood components. Springer, New York, pp 89–180 Esen A (1993) b-Glucosidases-biochemistry and molecular biology, ACS symposium series, 533. American Chemical Society, Washington, DC Foster RC (1988) Micro environment of Soil Microorganism. Biology and Fertility of Soils 6:189–203 Fitzgerald JW (1976) Sulphate ester formation and hydrolysis: a potentially important yet often ignored aspect of the sulphur cycle of aerobic soils. Bacteriol Rev 40:628–721 Frank T, Malkomes HP (1993) Influence of temperature on microbial activities and their reaction to the herbicide Goltix in different soils under laboratory conditions. Zentralblatt f€ur Mikrobiol 148:403–412 Ganeshamurthy AM, Singh G, Singh NT (1995) Sulphur status and response of rice to sulphur on some soils of Andaman and Nicobar Islands. J Indian Soc Soil Sci 43:637–641 Garcia C, Herna´ndez T (1997) Biological and biochemical indicators in derelict soils subject to erosion. Soil Biol Biochem 29:171–177 Giri V, Anuradha, Nandhini A, Geetha M, Gautham P (2004) A novel medium for the enhanced cell growth and production of prodigiosin from Serratia marcescens isolated from soil. BMC Microbiol 4:1–10 Giri B, Pham HG, Rina K, Ram P, Minu S, Garg A, Oelmuller R, Varma A (2005) Mycorrhizosphere: strategies and functions. In: Buscot F, Varma A (eds) Microorganisms in soils: roles in genesis and function, 3. Springer, Heidelberg, pp 213–252 Glinski J, Stepniewski W (1985) Soil aeration and its role for plants. CRC, Boca Raton, FL Gomah AM (1980) CM-cellulase activity in soil as affected by addition of organic material, temperature, storage and drying and wetting cycles. Zeitschrift fuer Pflanzenernaehrung und Bodenkunde 143:349–356 Gupta VVSR, Farrell RE, Germida JJ (1993) Activity of arylsuphatases in Saskatchewan soils. Can J Soil Sci 73:341–347 Haanstra L, Doelman P (1991) An ecological dose–response model approach to short- and longterm effects of heavy metals on arylsulphatase activity in soil. Biol Fertil Soils 1:18–23 Hans YW, Snivasan VR (1969) Purification and characterization of b-glucosidases of Alcaligenes faecalis. J Bacteriol 100:1355–1363 Hope CFA, Burns RG (1987) Activity, origins and location of cellulases in a silt loam soil. Biol Fert Soils 5:164–170 Izaguirre-Mayoral ML, Flores S, Carballo O (2002) Determination of acid phosphatases and dehydrogenase activities in the rhizosphere of nodulated legume species native to two contrasting savannah sites in Venezuela. Biol Fertil Soils 35:470–472 James ES, Russel LW, Mitrick A (1991) Phosphate stress response in hydroponically grown maize. Plant Soil 132:85–90 ¨ hlinger R, Kandeler E, Margesin R (eds) Methods in Kandeler E (1996) Nitrate. In: Schinner F, O soil biology. Springer, Berlin, pp 408–410 Kanfer JN, Mumford RA, Raghavan SS, Byrd J (1974) Purification of b-glucosidase activities from bovine spleen affinity chromatography. Anal Biochem 60:200–205 Khaziyev FK, Gulke AY (1991) Enzymatic activity of soils under agrocenosesa: status and problems. Pochvovedenie 8:88–103 King NJ (1967) Glucoamylase of Coniophora cerebella. Biochem J 105:577–583 Klein DA (1989) Cellulose functions in arid soil development. Arid Soil Res Rehab 3(1):85–198 Kranner I, Beckett RP, Varma AK (2002) Protocols in lichenology, culturing, biochemistry, ecophysiology and use in biomonitoring. Springer Lab Manual. Springer, Heidelberg

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Kuperman RG, Carreiro MM (1997) Soil heavy metal concentrations, microbial biomass and enzyme activities in a contaminated grassland ecosystem. Soil Biol Biochem 29:179–190 Ladd JN, Jackson RB (1982) In: Stevenson FJ (Ed) Nitrogen in agricultural soils. American Society of Agronomy, WI, pp 173–228 Leiro´s MC, Trasar-Cepeda C, Garcia´-Ferna´ndez F, Gil-Sotre´s F (1999) Defining the validity of a biochemical index of soil quality. Biol Fertil Soils 30:140–146 Liesack W, Stackebrandt E (1992) Occurrence of novel groups of the domain bacteria as revealed by analysis of genetic material isolated from an Australian Terrestrial Environment. J Bacteriol 174:5072–5078 Li Y, Guohua M, Fanjun C, Jianhua Z, Fusuo Z (2004) Rhizosphere effect and root growth of two maize (Zea mays L.) genotypes with contrasting P efficiency at low P availability. Plant Sci 167:217–223 Loper JE, Hack C, Schroth MN (1985) Population dynamics of soil Pseudomonads in the rhizosphere of potato. Appl Environ Microbiol 60:2394–2399 Lynch JM (1987a) Microbial interactions in the rhizosphere. Soil Microorg 30:33–41 Lynch JM (1987b) Soil biology-accomplishments and potential. Soil Sci Soc Am J 51:1409–1412 Madejo´n E, Burgos P, Lo´pez R, Cabrera F (2001) Soil enzymatic response to addition of heavy metals with organic residues. Biol Fertil Soils 34:144–150 Malla R, Tanaka Y, Mori K and Totawat KL (2007) Effect of Short-term Sewage Irrigation on Chemical Build Up in Soils and Vegetables. The Agricultural Engineering International: The CIGR Ejournal. Manuscript LW 07 006 Vol. IX Martinez CE, Tabatabai MA (1997) Decomposition of biotechnology byproducts in soils. J Environ Qual 26:625–632 Mayaudon J, Batistic L, Sarkar JM (1975) Propriete´s des activite´s proteolitiques extraites des sols frais. Soil Biol Biochem 7:281–286 McCarthy GW, Siddaramappa R, Reight RJ, Coddling EE, Gao G (1994) Evaluation of coal combustion byproducts as soil liming materials: their influence on soil pH and enzyme activities. Biol Fertil Soils 17:167–172 McLaren AD (1975) Soil as a system of humus and clay immobilized enzymes. Chem Scripta 8:97–99 Miwa T, Ceng CT, Fujisaki M, Toishi A (1937) Zur Frage der Spezifitat der Glykosidasen. I. Verhalted von b-d-glucosidases verschiedener Herkunft gegenuberden b-d-Glucosiden mit verschiedenen Aglykonen. Acta Phytochim (Tokyo) 10:155–170 Nannipieri P, Sequi P, Fusi P (1996) Humus and enzyme activity. In: Piccolo A (ed) Humic substances in terrestrial ecosystems. Elsevier, New York, pp 293–328 Ndakidemi PA (2006) Manipulating legume/cereal mixtures to optimize the above and below ground interactions in the traditional African cropping systems. Afr J Biotechnol 5:2526–2533 Ndiaye EL, Sandeno JM, McGrath D, Dick RP (2000) Integrative biological indicators for detecting change in soil quality. Am J Altern Agric 15:26–36 Paul EP, Clark FE (1989) Soil Microbiology and Biochemistry. Academic Press, Sandiego Pancholy SK, Rice EL (1973) Soil enzymes in relation to old field succession; amylase, cellulose, invertase, dehydrogenase and urease. Soil Sci Soc Am J 37:47–50 Patra DD, Brookes PC, Coleman K, Jenkinson DS (1990) Seasonal changes of soil microbial biomass in an arable and a grassland soil which have been under uniform management for many years. Soil Biol Biochem 22:739–742 Pazur JH (1965) Enzymes in the synthesis and hydrolysis of starch. In: Whistler R, Paschall EF (eds) Starch: chemistry and technology, vol 1, Fundamental aspects. Academic, New York, pp 133–175 Petker AS, Rai PK (1992) Effect of fungicides on activity, secretion of some extra cellular enzymes and growth of Alternaria alternata. Indian J Appl Pure Biol 7:57–59 Pettit NM, Smith ARJ, Freedman RB, Burns RG (1976) Soil urease: activity, stability and kinetic properties. Soil Biol Biochem 8:479–484

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Pham GH, Kumari R, Singh A, Sachdeva M, Prasad R, Kaldorf M, Buscot F, Oelmuller R, Tatjana P, Weiss M, Hampp R, Varma A (2004) Axenic culture of Piriformospora indica. In: Plant Surface Microbiology (eds A Varma, L Abbott, D Werner and R Hampp). Springer-Verlag, Germany 2004:593–616 Pitchel JR, Hayes JM (1990) Influence of fly ash on soil microbial activity and populations. J Environ Qual 19:593–597 Reddy GB, Faza A (1989) Dehydrogenase activity in sludge amended soil. Soil Biol Biochem 21:327 Richmond PA (1991) Occurrence and functions of native cellulose. In: Haigler CH, Weimer PJ (eds) Biosynthesis and biodegradation of cellulose. Dekker, New York, pp 5–23 Ross DJ (1968) Activities of enzymes hydrolysing sucrose and starch in some grassland soils. Trans 9th International Congress Soil-Science 3:299–308 Ross DJ (1975) Studies on a climosequence of soils in tussock grasslands-5. Invertase and amylase activities of topsoils and their relationships with other properties. NZ J Sci 18:511–518 Ross DJ (1976) Invertase and amylase activities in ryegrass and white clover plants and their relationships with activities in soils under pasture. Soil Biol Biochem 8:351–356 Ross DJ, Roberts HS (1970) Enzyme activities and oxygen uptakes of soils under pasture in temperature and rainfall sequences. J Soil Sci 21:368–381 Rotini OT (1935) La trasformazione enzimatica dell’urea nel terreno. Ann Labor Ric Ferm Spallanrani 3:143–154 Rubidge T (1977) The effect of moisture content and incubation temperature upon the potential cellulase activity of John Innes no. 1 soil (ISSN. 0020-6164). Int Biodeterior Bul 13:39–44 Schmidt G, Laskowski M Sr (1961) Phosphate ester cleavage (Survey). In: Boyer PD, Lardy H, Myrback K (eds) The enzymes, 2nd edn. Academic, New York, pp 3–35 Schussler A, Kluge M (2001) Geosiphon pyriforme, an endosymbiosis between fungus and cyanobacteria and its meaning as a model system for arbuscular mycorrhizal research. In: Esser K, Hock B (eds) The Mycota, 9. Springer, Heidelberg, pp 151–161 Shawale JG, Sadana J (1981) Purification, characterization and properties of b-glucosidase enzyme from Sclerotium rolfsii. Arch Biochem Biophys 207:185–196 Sinsabaugh RL, Antibus RK, Linkins AE (1991) An enzymic approach to the analysis of microbial activity during plant litter decomposition. Agric Ecosyst Environ 34:43–54 Sinsabaugh RL, Linkins AE (1989) Natural disturbance and the activity of Trichoderma viride cellulase complex. Soil Biol Biochem 21:835–839 Skujins J (1978) Soil enzymology and fertility index – a fallacy? History of abiotic soil enzyme research. In: Burns RG (ed) Soil enzymes. Academic, London, UK Speir TW, Ross DJ (1978) Soil phosphatase and sulphatase. In: Burns RG (ed) Soil enzymes. Academic, London, UK, pp 197–250 Staddon WJ, Duchesne LC, Trevors JT (1998) Impact of clear-cutting and prescribed burning on microbial diversity and community structure in a Jack pine (Pinus banksiana Lamb.) clear-cut using BiOLOG gram-negative microplates. World J Microbiol Technol 14:119–123 Stevenson FJ (1986) Cycles of Soil-Carbon, Nitrogen, Phosphorus, Sulfur, Micronutrients; Wiley InterScience Publ, John Wiley & Sons, New York Subba Rao NS (1997) Soil microbiology. IBH Pub, Oxford Tabatabai MA (1977) Effect of trace elements on urease activity in soils. Soil Biol Biochem 9:9–13 Tabatabai MA (1982) Soil enzyme. In: Page AL (ed) Methods of soil analysis, part 2. American Society of Agronomy, Madison, WI, pp 903–948 Tabatabai MA (1994) Soil enzymes. In: Weaver RW, Angle JS, Bottomley PS (eds) Methods of soil analysis, part 2. Microbiological and biochemical properties, SSSA book series no. 5. Soil Science Society America, Madison, WI, pp 775–833 Thoma JA, Spradlin JE, Dygert S (1971) Plant and animal amylases. In: Boyer PD (ed) The enzymes. International Society of Soil-Science, Netherlands 5:115–189

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Tisserant B, Gianinazzi-Pearson V, Gianinazzi S, Gollotte A (1993) In planta histochemical staining of fungal alkaline phosphatase activity for analysis of efficient arbuscular mycorrhizal infections. Mycol Res 97:245–250 Trevors JT (1984) Dehydrogenase activity in soil: a comparison between the INT and TTC assay. Soil Biol Biochem 16:673–674 Vincent PG, Sisler HD (1968) Mechanisms of antifungal action of 2,4,5,6-tetrachloroisopathalonitrile. Physiol Plant 21:1249–1264 Visscher PT, Vandenede FP, Schaub BEM, van Gemerden H (1992) Competition between anoxygenic phototropic bacteria and colorless sulfur bacteria in microbial mat. FEMS Microbiol Ecol 101:51–58 Wenzel WW, Pollak MA, Riedler C, Zischka RR, Blum WEH (1995) Influence of site conditions and heavy metals on enzyme activities of forest topsoil. In: Huang PM, Berthelin J, Bollag JM, McGill WN, Page AL (eds) Environmental impact of soil component interactions-metals, other inorganics, and microbial activities. CRC, Baton Rouge, LA, pp 211–225 White AR (1982) Visualization of cellulases and cellulose degradation. In: Brown RM Jr (ed) Cellulose and other natural polymer systems: biogenesis, structure, and degradation. Plenum, New York, pp 489–509 Wieland G, Neumann R, Backhaus H (2001) Variation of microbial communities in soil, rhizosphere, and rhizoplane in response to crop species, soil type, and crop development. Appl Environ Microbiol 67:5849–5854 Wilke BM (1991) Effect of single and successive additions of cadmium, nickel and zinc on carbon dioxide evolution and dehydrogenase activity in a sandy Luvisol. Biol Fertil Soils 11:34–37 Williams CH (1975) The chemical nature of sulphur compounds in soil. In: McLachlan KD (ed) Sulphur in Australasian agriculture. Sydney University Press, NSW, pp 21–30 Wright AL, Reddy KR (2001) Phosphorus loading effects on extracellular enzyme activity in Everglades wetland soil. Soil Sci Soc Am J 65:588–595 Yadav RS, Tarafdar JC (2001) Influence of organic and inorganic phosphorous supply on the maximum secretion of acid phosphatase by plants. Biol Fertil Soils 34:140–143 Yang Z, Liu S, Zheng D, Feng S (2006) Effects of cadium, zinc and lead on soil enzyme activities. J Environ Sci 18:1135–1141 Zantua MI, Bremner JM (1977) Stability of urease in soils. Soil Biol Biochem 9:135–140

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Chapter 2

Role of Enzymes in Maintaining Soil Health Shonkor Kumar Das and Ajit Varma

2.1

Introduction

Enzymes are the vital activators in life processes, likewise in the soil they are known to play a substantial role in maintaining soil health and its environment. The enzymatic activity in the soil is mainly of microbial origin, being derived from intracellular, cell-associated or free enzymes. A unique balance of chemical, physical, and biological (including microbial especially enzyme activities) components contribute to maintaining soil health. Evaluation of soil health therefore requires indicators of all these components. Healthy soils are essential for the integrity of terrestrial ecosystems to remain intact or to recover from disturbances, such as drought, climate change, pest infestation, pollution, and human exploitation including agriculture (Ellert et al. 1997). Deterioration of soil, and thereby soil health, is of concern for human, animal, and plant health because air, groundwater, and surface water consumed by humans, can be adversely affected by mismanaged and contaminated soil (Singer and Ewing 2000). As soil is the part of the terrestrial environment and supports all terrestrial life forms, protection of soil is therefore of high priority and a thorough understanding of soil enzymes activities is a critical factor in assuring that soil remains healthy. A better understanding of the role of this soil enzymes activity in maintaining the soil health will potentially provide a unique opportunity for an integrated biological assessment of soils due to their crucial role in several soil biological activities, their ease of measurement, and their rapid response to changes in soil management. Although there have been extensive studies on soil enzymes, little has been reported on their roles in maintaining soil health. Thus, it is authoritative to S.K. Das (*) Department of Applied Chemistry and Biotechnology, Graduate School of Engineering, University of Fukui, 3-9-1, Bunkyo, Fukui 910-8507, Japan e-mail: [email protected] A. Varma Amity Institute of Microbial Technology, Amity University, Sector 125, Noida 201303, Uttar Pradesh, India

G. Shukla and A. Varma (eds.), Soil Enzymology, Soil Biology 22, DOI 10.1007/978-3-642-14225-3_2, # Springer-Verlag Berlin Heidelberg 2011

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understand the roles of these enzymes’ activity and their efficiency to maintain soil health for future betterment of soil research and soil biology.

2.2

Soil Enzymes

Soil enzymes are a group of enzymes whose usual inhabitants are the soil and are continuously playing an important role in maintaining soil ecology, physical and chemical properties, fertility, and soil health. These enzymes play key biochemical functions in the overall process of organic matter decomposition in the soil system (Sinsabaugh et al. 1991). They are important in catalyzing several vital reactions necessary for the life processes of micro-organisms in soils and the stabilization of soil structure, the decomposition of organic wastes, organic matter formation, and nutrient cycling, hence playing an important role in agriculture (Dick et al. 1994; Dick 1997). All soils contain a group of enzymes that determine soil metabolic processes (McLaren 1975) which, in turn, depend on its physical, chemical, microbiological, and biochemical properties. The enzyme levels in soil systems vary in amounts primarily due to the fact that each soil type has different amounts of organic matter content, composition, and activity of its living organisms and intensity of biological processes. In practice, the biochemical reactions are brought about largely through the catalytic contribution of enzymes and variable substrates that serve as energy sources for microorganisms (Kiss et al. 1978). These enzymes may include amylase, arylsulphatases, b-glucosidase, cellulose, chitinase, dehydrogenase, phosphatase, protease, and urease released from plants (Miwa et al. 1937), animals (Kanfer et al. 1974), organic compounds, and microorganisms (James et al. 1991; Richmond 1991; Shawale and Sadana 1981) and soils (Gupta et al. 1993; Ganeshamurthy et al. 1995).

2.2.1

Kind of Soil Enzymes

2.2.1.1

Constitutive

Always present in nearly constant amounts in a cell (not affected by addition of any particular substrate – genes always expressed). (Pyrophosphatase)

2.2.1.2

Inducible

Present only in trace amounts or not at all, but quickly increases in concentration when its substrate is present. (Amidase)

Both types of enzymes are present in the soil.

2 Role of Enzymes in Maintaining Soil Health

2.2.2

27

Origin and State of Soil Enzymes

Although the general origins of soil enzymes are (a) microorganisms-living and dead, (b) plant roots and plant residues and, (c) soil animals; the state of soil enzymes in the soil is different as below State-1: Role of clays Most activity associated with clays Increased resistance to proteolysis and microbial attack Increases the temperature of inactivation State-2: Role of organic matter Humus material provides stability to soil nitrogen compounds Enzymes attached to insoluble organic matrices exhibit pH and temperature changes Inability to purify soil enzymes free of soil organic matters (bound to organic matter) State-3: Role of clay–organic matter complexes Lignin + bentonite (clay) protect enzymes against proteolitic attack, but not bentonite alone Enzymes are bound to organic matter which is then bound to clay

2.2.3

Importance of Soil Enzymes

Release of nutrients into the soil by means of organic matter degradation Identification of soils Identification of microbial activity Importance of soil enzymes as sensitive indicators of ecological change

2.2.4

Application of Soil Enzymes

Correlation with soil fertility Correlation with microbial activity Correlation with biochemical cycling of various elements in soil (C, N, S) Degree of pollution (heavy metals, SO4) To assess the successional stages of an ecosystem Forensic purposes Rapid degradation of pesticides Disease studies Enzyme activity in soil fluctuates with environment.

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2.3 2.3.1

S.K. Das and A. Varma

Soil Health Definition

The definition of soil health must be broad enough to encompass the many functions of soil, e.g., environmental filter, plant growth, and water regulation (Doran and Safley 1997). Definitions of air and water quality standards have existed for a long time, while a similar definition does not exist for soil. A definition of soil health based on this concept would encompass only a small fraction of the many roles soil play (Singh et al. 1999). Soil health is the net result of on-going conservation and degradation processes, depending highly on the biological component of the soil ecosystem, and influences plant health, environmental health, food safety, and quality (Halvorson et al. 1997; Parr et al. 1992). Several definitions of soil health have been proposed during the last decades. Historically, the term soil quality described the status of soil as related to agricultural productivity or fertility (Singer et al. 1999). In the 1990s, it was proposed that soil quality was not limited to soil productivity but instead expanded to encompass interactions with the surrounding environment, including the implications for human and animal health. In this regard, several examples of definitions of soil quality have been suggested (Doran and Parkin 1994). In the mid-1990s, the term soil health was introduced. For example, a program to assess and monitor soil health in Canada used the terms quality and health synonymously to describe the ability of soil to support crop growth without becoming degraded or otherwise harming the environment (Acton and Gregorich 1995). Others broadened the definition of soil health to capture the ecological attributes of soil, and went beyond its capacity to simply produce particular crops. These attributes are chiefly associated with biodiversity, food web structure, and functional measures (Pankhurst et al. 1997). Several numbers have been recognized for soil health which are as follows: 1. The continued capacity of soil to function as a vital living system, within the ecosystem and land-use boundaries, to sustain biological productivity, promote the quality of air and water environments, and maintain plant, animal, and human health (Doran and Safley 1997). 2. Soil health is an assessment of ability of a soil to meet its range of ecosystem functions as appropriate to its environment. 3. Soil health can also be defined as the continued capacity of a specific kind of soil to function as a vital living system, within natural or managed ecosystem boundaries, to sustain plant and animal productivity, to maintain or enhance the quality of air and water environments, and to support human health and habitation.

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2.3.2

29

Aspects of Soil Health

The term soil health is used to assess the ability of a soil to Sustain plant and animal productivity and diversity Maintain or enhance water and air quality Support human health and habitation The underlying principle in the use of the term “soil health” is that soil is not just a growing medium; rather, it is a living, dynamic and ever-so-subtly changing environment. We can use the human health analogy and categorize a healthy soil as one In a state of composite well-being in terms of biological, chemical, and physical properties Not diseased or infirmed (i.e., not degraded, nor degrading), nor causing negative off-site impacts With each of its qualities cooperatively functioning such that the soil reaches its full potential and resists degradation Providing a full range of functions (especially nutrient, carbon, and water cycling), and in such a way that it maintains this capacity into the future.

2.3.3

Interpretation of Soil Health

Different soils will have different benchmarks of health depending on the “inherited” qualities, and on the geographic circumstance of the soil. The generic aspects defining a healthy soil can be considered as follows “Productive” options are broad Life diversity is broad Absorbency, storing, recycling, and processing is high in relation to limits set by climate Water runoff quality is of high standard Low entropy No damage to or loss of the fundamental components This translates to A comprehensive cover of vegetation Carbon levels relatively close to the limits set by soil type and climate Little leakage of nutrients from the ecosystem

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Biological productivity relatively close to the limits set by the soil environment and climate Only geological rates of erosion No accumulation of contaminants The ecosystem does not rely excessively on inputs of fossil energy An unhealthy soil thus is the simple converse of the above.

2.3.4

Pressures on Soil Health Towards Impacts

The flow chart given below is a simple description of soil health factors and their impacts (Fig. 2.1).

Pressures on soil health Climate, natural events, urbanization, agriculture, forestry, waste disposal etc.

Soil Health

Atmospheric balance

Human health

Animal health

Plant health

Soil ecosystem health

Soil microbial community health

Leaching to groundwater Surface run-off

End points Fig. 2.1 Policy relevant end points of soil health monitoring

2 Role of Enzymes in Maintaining Soil Health

2.4 2.4.1

31

Indicators of Soil Health Microorganism as Indicators of Soil Health

The biological activity in soil is largely concentrated in the topsoil, the depth of which may vary from a few to 30 cm. In the topsoil, the biological components occupy a tiny fraction (<0.5%) of the total soil volume and make up less than 10% of the total organic matter in the soil. These biological components consist mainly of soil organisms, especially microorganisms. Despite of their small volume in soil, microorganisms are key players in the cycling of nitrogen, sulfur, and phosphorus, and the decomposition of organic residues. Thereby they affect nutrient and carbon cycling on a global scale (Pankhurst et al. 1997). That is, the energy input into the soil ecosystems is derived from the microbial decomposition of dead plant and animal organic matter. The organic residues are, in this way, converted to biomass or mineralized to CO2, H2O, mineral nitrogen, phosphorus, and other nutrients (Bloem et al. 1997). Microorganisms are further associated with the transformation and degradation of waste materials and synthetic organic compounds (Torstensson et al. 1998). In addition to the effect on nutrient cycling, microorganisms also affect the physical properties of the soil. Production of extra-cellular polysaccharides and other cellular debris by microorganisms help in maintaining soil structure as well as soil health. Thereby, they also affect water holding capacity, infiltration rate, crusting, erodibility, and susceptibility to compaction (Elliott et al. 1996). Microorganisms possess the ability to give an integrated measure of soil health, an aspect that cannot be obtained with physical/chemical measures and/or analyses of diversity of higher organisms. Microorganisms respond quickly to changes; hence they rapidly adapt to environmental conditions, and thus they can be used for soil health assessment, and changes in microbial populations and activities may therefore function as an excellent indicator of change in soil health (Kennedy and Papendick 1995; Pankhurst et al. 1995). Microorganisms also respond quickly to environmental stress compared to higher organisms, as they have intimate relations with their surroundings due to their high surface to volume ratio. In some instances, changes in microbial populations or activity can precede detectable changes in the soil’s physical and chemical properties, thereby providing an early sign of soil improvement or an early warning of soil degradation (Pankhurst et al. 1995). The impact of some chemicals on soil health is dependent on microbial activities. For example, the concentration of heavy metals in soil will not change over small time periods, but their bioavailability may. In this way, soil enzymes are acting as important indicators of soil.

2.4.2

Soil Enzymes as Indicators of Soil Health

Enzymes are the direct mediators for biological catabolism of soil organic and mineral components. Thus, these catalysts provide a meaningful assessment of

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Table 2.1 Soil enzymes as indicators of soil health Soil enzyme Enzyme reaction Dehydrogenase Electron transport system b-glucosidase Cellobiose hydrolysis Cellulase Cellulose hydrolysis Phenol oxidase Lignin hydrolysis Urease Urea hydrolysis Amidase N-mineralization Phosphatase Release of PO4 Arylsulphatase Release of SO4 Soil enzymes Hydrolysis

Indicator of microbial activity C-cycling C-cycling C-cycling C-cycling N-cycling N-cycling P-cycling S-cycling General organic matter degradative enzyme activities

reaction rates for important soil processes. Soil enzyme activities (1) are often closely related to soil organic matter, soil physical properties and microbial activity or biomass, (2) changes much sooner than other parameters, thus providing early indications of changes in soil health, and (3) involve simple procedures (Dick et al. 1996). In addition, soil enzyme activities can be used as measures of microbial activity, soil productivity, and inhibiting effects of pollutants (Tate 1995). Easy, well-documented assays are available for a large number of soil enzyme activities (Dick et al. 1996; Tabatabai 1994a, b). These include dehydrogenase, glucosidases, urease, amidases, phosphatases, arylsulphatase, cellulases, and phenol oxidases as shown in Table 2.1.

2.5

Potential Roles of Soil Enzymes in Maintaining Soil Health

A number of soil enzymes and their respective roles in maintaining soil health are stated below

2.5.1

Amylase

The starch hydrolyzing enzyme amylase (Ross 1976) is known to be constituted by a-amylase and b-amylase (King 1967; Thoma et al. 1971). The a-amylases are synthesized by plants, animals, and microorganisms, whereas, b-amylase is synthesized mainly by plants (Pazur 1965; Thoma et al. 1971). This enzyme is widely distributed in plants and soils so it plays a significant role in the breakdown of starch, which converts starch like substrates to glucose and/or oligosaccharides and b-amylase, which converts starch to maltose (Thoma et al. 1971). Studies have, however, indicated that the roles and activities of a-amylase and b-amylase enzymes may be influenced by different factors ranging from cultural practices, type of vegetation, environment and soil types (Pancholy and Rice 1973; Ross 1975). For example, plants may influence the amylase enzyme activities of soil by

2 Role of Enzymes in Maintaining Soil Health

33

directly supplying enzymes from their residues or excreted compounds, or indirectly providing substrates for the synthetic activities of microorganisms. Greater understanding is required of the significance of these enzymes in the soil, and to enable proper management techniques to be devised to maximize the benefits that may be derived from such enzymes.

2.5.2

Arylsulphatases

This is due to the fact that certain proportions of sulphur in different soil profiles are bound into organic compounds and are indirectly available to plants. Arylsulphatases are typically widespread in nature (Dodgson et al. 1982) as well as in soils (Gupta et al. 1993; Ganeshamurthy et al. 1995). They are responsible for the hydrolysis of sulphate esters in the soil (Kertesz and Mirleau 2004) and are secreted by bacteria into the external environment as a response to sulphur limitation (McGill and Colle 1981). Its occurrence in different soil systems is often correlated with microbial biomass and rate of S immobilization (Klose and Tabatabai 1999; Vong et al. 2003). This enzyme has a role in the hydrolysis of aromatic sulphate esters (R–O–SO3 ) to phenols (R–OH) and sulfate, or sulfate sulfur (SO4 2 or SO4–S) (Tabatabai 1994a, b). Soil is affected by various environmental factors (Burns 1982) such as heavy metal pollution (Tyler 1981); pH changes in the soil solution (Acosta-Martı´nez and Tabatabai 2000); organic matter content and its type (Sarathchandra and Perrott 1981); such as absorption to particles surfaces in soils, and the activity persistence of extra cellular arylsulfatases in the soil. Considering the importance of S in plant nutrition, a better understanding of the role(s) of arylsulfatases in S mobilization in agricultural soils is critical. So far, very little is known about specific microbial genera or species that play an important role in the soil organosulphur circle (Kertesz and Mirleau 2004) in which arylsulphatases is the key enzyme.

2.5.3

b-Glucosidase

Glucosidase is a common and predominant enzyme in soils (Eivazi and Tabatabai 1988; Tabatabai 1994a, b). It is named according to the type of bond that it hydrolyses. This enzyme plays an important role in soils because it is involved in catalyzing the hydrolysis and biodegradation of various b-glucosidase present in plant debris decomposing in the ecosystem (Ajwa and Tabatabai 1994; Martinez and Tabatabai 1997). Its final product is glucose, an important C energy source of life to microbes in the soil (Esen 1993). b-glucosidase is characteristically useful as a soil quality indicator, and may give a reflection of past biological activity, the capacity of soil to stabilize the soil organic matter, and can be used to detect management effect on soils (Bandick and Dick 1999; Ndiaye et al. 2000). This has greatly facilitated its

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adoption for soil quality testing (Bandick and Dick 1999). Some of the aglycons are known to be the precursors of the toxic substances, which cause soil sickness where plants are grown as monocrops (Patrick 1955; Borner 1958). b-Glucosidase enzyme is very sensitive to changes in pH, and soil management practices (Acosta-Martı´nez and Tabatabai 2000; Madejo´n et al. 2001). AcostaMartı´nez and Tabatabai 2000 reported b-glucosidase as sensitive to pH changes. This property can be used as a good biochemical indicator for measuring ecological changes resulting from soil acidification in situations involving activities of this enzyme. Consequently, more understanding of the b-glucosidase enzyme activities and factors influencing them in the ecosystem may contribute significantly to soil health studies.

2.5.4

Cellulases

Cellulose is the most abundant organic compound in the biosphere, comprising almost 50% of the biomass synthesized by photosynthetic fixation of CO2 (Eriksson et al. 1990). Growth and survival of microorganisms important in most agricultural soils depends on the carbon source contained in the cellulose occurring in the soils (Deng and Tabatabai 1994). However, for carbon to be released as an energy source for use by the microorganisms, cellulose in plant debris has to be degraded into glucose, cellobiose and high molecular weight oligosaccharides by cellulases enzymes (White 1982). Cellulases are a group of enzymes that catalyze the degradation of cellulose, polysaccharides built up of b-1,4 linked glucose units (Deng and Tabatabai 1994). It has been reported that cellulases in soils are derived mainly from plant debris incorporated into the soil, and that a limited amount may also originate from fungi and bacteria in soils (Richmond 1991). Demonstrating the effects of increasing concentrations of fungicides on cellulases activities, Petker and Rai (1992) showed that there was a decreasing effect with fungicides captan, cosan, thiram, zinels, and sandolex. More recently, Arinze and Yubedee (2000) reported that fungicides benlate, calixin, and captan inhibited cellulase activity in Fusarium monoliforme isolates. Captatol inhibited cellulose activity in the sandy loam soil (Atlas et al. 1978), and chlorothalonil showed a clear reduction in cellulase activity under flooded or non-flooded conditions (Vincent and Sisler 1968). Studies have shown that activities of cellulases in agricultural soils are affected by several factors. These include temperature, soil pH, water and oxygen contents (abiotic conditions), the chemical structure of organic matter and its location in the soil profile horizon (Deng and Tabatabai 1994; Alf and Nannipieri 1995), quality of organic matter/plant debris and soil mineral elements (Sinsabaugh and Linkins 1989; Deng and Tabatabai 1994) and the trace elements from fungicides (Deng and Tabatabai 1994; Arinze and Yubedee 2000). Srinivasulu and Rangaswamy 2006 reported a significantly more stimulatory effect of cellulases in black soil than red soil. For instance, chitin in the presence of cellulose induces the synthesis of chitinase and other cell wall lytic enzymes which promote the

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release of the intramural b-glucosidase into the medium. All these findings suggest that activities of cellulases can be used to give preliminary indication of some of the physical chemical properties of soil, thus, easing agricultural soil management strategies. Since cellulases enzymes play an important role in global recycling of the most abundant polymer, cellulose in nature, it would be of critical importance to understand this enzyme better so that it may be used more regularly as a predictive tool in our soil fertility programs. More information on the role of this enzyme is needed since it is affected by different factors, which may jeopardize its involvement in the decomposition of cellulolytic materials in the soil for microbial use and improved soil health in agricultural ecosystems.

2.5.5

Chitinase

Chitinase or chitinolytic enzymes are key enzymes responsible for the degradation and hydrolysis of chitin (polyb-1-4-(2-ncetamido-2-deoxy)-D-glucoside). They are also considered as the major structural component of many fungal cell walls that use the hyperparasitism mechanisms against pests/pathogen attack (Chet and Henis 1975; Chet 1987). These biological agents also reduce disease-producing agents by using other mechanisms such as antibiosis or competition mechanisms (Park 1960). This agriculturally important enzyme is produced or released by various organisms including plants and microorganisms (Deshpande 1986). Its presence in different forms in the ecosystem has demonstrated its effectiveness in the control of soil-borne diseases such as Sclerotium rolfsii and Rhizoctonia solani in beans and cotton, respectively (Ordentlich et al. 1988; Shapira et al. 1989). One of the mechanisms proposed involves lytic enzymes chitinase that cause the degradation of cell walls of pathogenic fungi (Ordentlich et al. 1988; Chet et al. 1990; Singh et al. 1999). As for its role in biological control of pests, moreover, due to environmental friendliness, there are so many avenues for the application of this enzyme for maintaining soil health and consequently, increase plant growth and final yields.

2.5.6

Dehydrogenase

The dehydrogenase enzyme activity is commonly used as an indicator of biological activity in soils (Burns 1978). This enzyme is considered to exist as an integral part of intact cells but does not accumulate extracellularly in the soil. Dehydrogenase enzyme is known to oxidize soil organic matter by transferring protons and electrons from substrates to acceptors. These processes are the part of respiration pathways of soil microorganisms and are closely related to the type of soil and soil air-water conditions (Kandeler 1996; Glinski and Stepniewski 1985). Since these processes are the part of respiration pathways of soil microorganisms, studies on the activities of dehydrogenase enzyme in the soil is very important as it may give

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indications of the potential of the soil to support biochemical processes which are essential for maintaining soil fertility as well as soil health. A study by Brzezinska et al. (1998) suggested that soil water content and temperature influence dehydrogenase activity indirectly by affecting the soil redox status. After flooding the soil, the oxygen present is rapidly exhausted so that a shift of the activity from aerobic to anaerobic microorganisms takes place. Such redox transformations are closely connected with respiration activity of soil microorganisms. They may serve as indicators of the microbiological redox systems in soils and can be considered a possible measure of microbial oxidative activity (Tabatabai 1982; Trevors 1984). For instance, lack of oxygen may trigger facultative anaerobes to initiate metabolic processes involving dehydrogenase activities and the use of Fe (III) forms as terminal electron acceptors (Bromfield 1954; Galstian and Awungian 1974), a process that may affect iron availability to plants in the ecosystem (Benckiser et al. 1984). Additionally, dehydrogenase enzyme is often used as a measure of any disruption caused by pesticides, trace elements or management practices to the soil (Reddy and Faza 1989; Wilke 1991; Frank and Malkomes 1993), as well as a direct measure of soil microbial activity (Trevors 1984; Garcia and Herna´ndez 1997). It can also indicate the type and significance of pollution in soils. For example, dehydrogenase enzyme is high in soils polluted with pulp and paper mill effluents (McCarthy et al. 1994) but low in soils polluted with fly ash (Pitchel and Hayes 1990). Similarly, higher activities of dehydrogenases have been reported at low doses of pesticides, and lower activities of the enzyme at higher doses of pesticides (Baruah and Mishra 1986).

2.5.7

Phosphatases

In soil ecosystems, these enzymes are believed to play critical roles in P cycles (Speir and Ross 1978) as evidence shows that they are correlated to P stress and plant growth. Apart from being good indicators of soil fertility, phosphatase enzyme plays a key role in the soil system (Eivazi and Tabatabai 1977; Dick et al. 2000). For example, when there is a signal indicating P deficiency in the soil, acid phosphatase secretion from plant roots is increased to enhance the solubilization and remobilization of phosphate, thus influencing the ability of the plant to cope with P-stressed conditions (Karthikeyan et al. 2002; Mudge et al. 2002; Versaw and Harrison 2002). Understanding the dynamics of enzyme activities in these systems is crucial for predicting their interactions as their activities may, in turn, regulate nutrient uptake and plant growth, later on, where soil health is concerned.

2.5.8

Proteases

Proteases in the soil play a significant role in N mineralization (Ladd and Jackson 1982), an important process regulating the amount of plant available N and plant

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growth. This enzyme in the soil is generally associated with inorganic and organic colloids (Burns 1982; Nannipieri et al. 1996). The amount of this extra cellular enzyme activity may be indicative not only of the biological capacity of soil for the enzymatic conversion of the substrate, which is independent of the extent of microbial activity, but might also have an important role in the ecology of microorganisms in the ecosystem (Burns 1982). There is a need to study the properties and factors affecting naturally occurring enzyme complexes such as those involving protease enzymes in the soil ecosystem as they may reveal some unknown role(s) in maintaining soil health and fertility.

2.5.9

Urease

Urease enzyme is responsible for the hydrolysis of urea fertilizers applied to the soil into NH3 and CO2 with the concomitant rise in soil pH (Andrews et al. 1989; Byrnes and Amberger 1989). This, in turn, results in a rapid N loss to the atmosphere through NH3 volatilization (Simpson et al. 1984; Simpson and Freney 1988). Due to this role, urease activities in soils have received a lot of attention since it was first reported by Rotini (1935), a process considered vital in the regulation of N supply to plants after urea fertilization. Soil urease originates mainly from plants (Polacco 1977) and microorganisms found as both intra- and extra-cellular enzymes (Burns 1986; Mobley and Hausinger 1989). On the other hand, urease extracted from plants or microorganisms is rapidly degraded in soil by proteolytic enzymes (Pettit et al. 1976; Zantua and Bremner 1977). This suggests that a significant fraction of ureolytic activity in the soil is carried out by extracellular urease, which is stabilized by immobilization on organic and mineral soil colloids. Urease activity in soils is influenced by many factors. These include cropping history, organic matter content of the soil, soil depth, soil amendments, heavy metals, and environmental factors such as temperatures (Tabatabai 1977; Yang et al. 2006). For example, studies have shown that urease was very sensitive to toxic concentrations of heavy metals (Yang et al. 2006). Generally, urease activity increases with increasing temperature. It is suggested that higher temperatures increase the activity coefficient of this enzyme. Therefore, it is recommended that urea be applied at times of the day when temperatures are low. Since urease plays a vital role in the hydrolysis of urea fertilizer, it is important to uncover other unknown factors that may reduce the efficiency of this enzyme in the ecosystem.

2.6

Conclusion

It is very essential to understand the possible roles of soil enzymes in order to maintain soil health and its fertility management in ecosystems. These enzymes, usually found in the soil, may have significant effects on soil biology,

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environmental management, growth and nutrient uptake in plants growing in ecosystems. Their activities may, however, be influenced by unknown cultural management practices either in a major or minor amount. Studies focusing the discovery of new enzymes from microbial diversity in the soil might be the most suitable practices that may positively influence their activities for improved plant growth as well as rendering the friendly biological environments in order to sustain other living beings.

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Chapter 3

Agricultural and Ecological Significance of Soil Enzymes: Soil Carbon Sequestration and Nutrient Cycling Wei Shi

3.1

Introduction

Organic matter decomposition is an important control of soil carbon sequestration and nutrient cycling. Due to sizable and complex nature of organic matter, decomposition often initiates with soil enzyme-catalyzed biochemical reactions. It is generally accepted that enzymatic depolymerization is a rate limiting step of decomposition. Therefore, research efforts are increasingly given to mechanisms that emphasize microbial enzyme production and soil enzyme activities that control carbon and nutrient dynamics. Agricultural and ecological significance of soil enzymes have been progressively expanded since the first report on soil enzymes about a century ago. Soil enzymes, once used as descriptive parameters, have now been appreciated for their multifaceted functions in microbial activities, soil processes, and ecosystem responses to management and global environmental change (Sinsabaugh 1994; Waldrop et al. 2004; Fenner et al. 2005a; Lipson et al. 2005; Finzi et al. 2006). Decomposition models with the component of soil enzymes appear to be robust not only in addressing fundamental questions, such as carbon and nutrient limitations on microbial activity, but also in predicting the degree of decomposition in relation to organic carbon chemistry and nutrient availability and under specific environmental conditions (Schimel and Weintraub 2003; Moorhead and Sinsabaugh 2006). While numerous enzymes exist in a given soil, their relative importance in terms of specific soil function and/or ecosystem service may vary. For example, dehydrogenases are often used as the indicator of soil microbial activity and lignocellulases have been recognized as essential soil enzymes to control the decomposition of soil organic matter and plant litters (Sinsabaugh et al. 1992; Sinsabaugh 1994).

W. Shi Department of Soil Science, North Carolina State University, Raleigh, NC 27695-7619, USA e-mail: [email protected]

G. Shukla and A. Varma (eds.), Soil Enzymology, Soil Biology 22, DOI 10.1007/978-3-642-14225-3_3, # Springer-Verlag Berlin Heidelberg 2011

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This chapter aims to discuss soil enzymes that can be used to learn about soil carbon sequestration and nutrient cycling. Certainly, it is not my intention to provide an exhaustive list of soil enzymes that are responsible for organic matter decomposition and nutrient mineralization or detail impacts of environmental factors, such as temperature, water availability, and soil pH on the activity of soil enzymes. Instead, the focus of this chapter is on soil enzymes that have been recognized as the proximate control of decomposition, principles by which a microbial community generates extracellular enzymes, and implications that are relevant to ecosystem responses to management and global environmental change.

3.2

3.2.1

Soil Enzymes for Organic Matter Decomposition and Nutrient Cycling Soil Enzymes of Primary Interest

Soil enzymes represent the enzymes that accumulate in soil and function extracellularly. Normally, a soil enzyme is associated with various abiotic and biotic soil constituents, such as soil aqueous phase, clay minerals, humic materials, cell debris, and viable but non-proliferating cells and, accordingly, its activity represents the sum of activities from these individual components (Ladd 1978; Burns 1982). While hundreds of extracellular enzymes may exist in a soil, only some has been quite often used to assess soil carbon and nutrient dynamics. Ligninase, cellulase, chitinase, and phosphatase are among extensively-exploited enzymes that catalyze organic matter decomposition and nutrient mineralization. Ligninase is a group of oxidative enzymes that modify phenolic-containing organics including polyphenols, lignin, and humus. As a typical ligninase, for example, soil phenol oxidase acts upon phenolics of various complexities by catalyzing the release of oxygen radicals (Hammel 1997; Claus 2004). Through this modification, simple phenolics may be fully degraded and complicated ones are perhaps partially oxidized, thereby generating phenolic intermediates (Burke and Cairney 2002; Claus 2004; Toberman et al. 2008b). Phenol oxidase activity can be limited by low oxygen pressure, low temperature, and acidic pH (McLatchey and Reddy 1998; Criquet et al. 1999; Sinsabaugh et al. 2008). A number of microbial species from fungi, bacteria, and actinomycetes are able to produce phenol oxidase (Hammel 1997; Hullo et al. 2001; Burke and Cairney 2002; Endo et al. 2003; Fenner et al. 2005b). Cellulase is a group of hydrolytic enzymes that catalyze the breakdown of glycosidic bonds in cellulose. Complete degradation of cellulose requires at least three enzymes: endo-b-1,4-glucanase, exo-b-1,4-glucanase, and b-glucosidase. Endoglucanase disrupts crystalline structure of cellulose via random cleavage of internal bonds; exoglucanase removes oligosaccharides, such as cellobiose and tetrasaccharides from the non-reducing ends of carbohydrate chains generated by

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endoglucanase; and glucosidase releases glucose from oligosaccharides (Alef and Nannipieri 1995). Several analytical methods have been developed for estimating the potential activity of soil endoglucanase, exoglucanase, glucosidase or their combination. As the major component of exoskeleton of insects and cell walls of fungi, chitin represents one of dominant forms of organic nitrogen in soil (Paul and Clark 1996). Chitinase is an essential class of enzymes that hydrolyze glycosidic bonds in chitin and release acetyl-glucosamine. Three enzymes, chitinase, chitobiase, and N-acetyl-bglucosaminidase act synergistically for the complete degradation of chitin. N-acetylb-glucosaminidase is often used as the indicator for the activity of soil chitinase. Phosphatase characterizes a broad group of enzymes that hydrolyze phosphate ester and subsequently release phosphate. In general, phosphatase has low substrate specificity and thus can act upon a number of different structurally-related substrates (Alef et al. 1995). Extensively studied soil phosphatase includes acid phosphatase and alkaline phosphatase that are mainly found in acid and alkaline soils, respectively (Tabatabai 1994).

3.2.2

Lignocellulolytic Enzymes as Predictors of Organic Matter Decomposition

A relationship between soil enzyme activity and decomposition rate is highly expected given that extracellular enzyme-catalyzed depolymerization, is the rate limiting step of decomposition. Lignin- and cellulose-degrading enzymes appear to be favorably chosen for predicting decomposition, mainly because soil organic matter and plant materials are of carbohydrate- and phenolic-based structures. Sinsabaugh et al. (1992) developed a regression model to estimate the mass loss rate of birch sticks using five lignocellulose-degrading enzymes, including b-glucosidase, endocellulase, exocellulase, xylosidase, and phenol oxidase. Enzyme activities accounted fairly well for variations in mass loss rate across upland, riparian, and lotic systems, with r2 ranging from 0.65 for endocellulase to 0.83 for b-glucosidase. Furthermore, the predictability of soil enzymes was improved by integrating activities of the five enzymes into a single and simple variable, referred to lignocellulase activity which explained 83% of mass loss variation. In a later study, an integrated cellulase activity was also used to estimate the decomposition rate of litters of various deciduous species (Sinsabaugh and Linkins 1993). Lignocellulolytic enzymes as predictors of organic matter decomposition appear to be supported by a number of nitrogen fertilization experiments, where changes in decomposition rate of plant litters are found to be positively associated with changes in cellulase and/or phenol oxidase activities (Carreiro et al. 2000; Saiya-Cork et al. 2002; DeForest et al. 2004; Waldrop et al. 2004; Sinsabaugh et al. 2005). An oxidative enzyme-based “latch” mechanism has been proposed to elucidate soil organic carbon retention in peatlands (Freeman et al. 2001a). Under limited

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oxygen supply, low phenol oxidase activity slows down degradation of phenolic compounds in soil aqueous phase. Because of phenolic toxicity to hydrolytic enzymes (Pind et al. 1994; Freeman et al. 2001a), phenolic accumulation in soil solution can inhibit hydrolytic degradation of organic matter and therefore benefit soil carbon sequestration. Soil oxidative enzymes have also been considered as the proximate control for organic matter decomposition, in other ecosystems. Differences in soil carbon storage among various habitats appear to reflect differences in the activities of oxidative enzymes. Ecosystems in high and low latitudes both have a low rate of primary production, but their soil organic carbon contents differ considerably. In low latitude ecosystems where phenol oxidase and peroxidase are high, soil organic carbon is low. In contrast, great soil organic carbon exists in high altitude ecosystems where the activities of phenol oxidase and peroxidase are constrained by low pH, low temperature, and low oxygen availability (Sinsabaugh et al. 2008).

3.2.3

Enzymes as Indicators of Soil Nutrient Availability

Soil enzymes seem to be informative to capture soil nutrient supplying capacity. Positive correlations between the activity of soil enzymes and nutrient mineralization have been reported in agricultural soils. In long-term nirogen-fertilization cropping systems, for example, the activity of N-acetyl-glucosaminidase was tightly assoicated with nitrogen mineralization potential (Ekenler and Tabatabai 2002). Nitrogen minerlization from a broad range of soils in the North Center region of the United States was well correlated with the activities of N-acetyl-glucosaminidase and arylamidase (Ekenler and Tabatabai 2004; Dodor and Tabatai 2007). Enzyme activity can also be used to indicate soil nutrient availability. A number of studies reveal that activities of soil phosphorus- and nitrogen-mineralization enzymes are negatively related to soil available phosphorous and nitrogen, respectively (Speir and Ross 1978; Sinsabaugh et al. 1993; Kang and Freeman 1999). In various cropping systems in the semi-arid region of Oregon, USA, for example, amidase and urease activitites have been found to be inversely related to application rates of synthetic nitrogen fertilizers (Dick 1992; Bandick and Dick 1999). Allison et al. (2008) also reported that nitrogen fertilization significantly suppressed the activities of protein- and chitin-degrading enzymes.

3.3

3.3.1

Microbial Enzyme Production as Affected by Organic Carbon and Nutrient Availability Positive and Negative Feedback Mechanisms

During evolution, any type of microbes may have developed some cost-efficient strategy to explore resource. Microbes produce extracellular enzymes perhaps only

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when the enzymes can help to better use the resource and therefore lead to optimal microbial growth and metabolisms. This cost-efficient strategy predicts that microbial enzyme production can be low when the end products of enzymatic reactions, e.g., nutrients, are abundant or when the substrates of enzymatic reactions, e.g., complex organics, are limited (Allison and Vitousek 2005). Soil enzyme activities are expected to be suppressed by the addition of nutrients or induced by the addition of complex organics. End product suppression on enzymes appears to be supported by numerous observations that the activity of a nutrient-mineralization enzyme is inversely related to the nutrient availability. Olander and Vitousek (2000) demonstrated that negative feedback was indeed the cause for the reduction of phosphatase and chitinase activities after long-term respective phosphorus and nitrogen fertilization. They argued that there were no direct and adverse effects of fertilization on enzyme activity because phosphatase and chitinase did not respond to short-term fertilization. Inverse relationship between phosphorus availability and phosphatase activity seems to be more common than the one between nitrogen availability and nitrogenmineralization enzymes. This is because organic phosphorus largely exists in the form of phosphate esters and is mineralized mainly through phosphatase catalysis. In contrast, organic nitrogen appears in various forms and can also be tightly associated with organic carbon. Not only do diverse nitrogen-mineralization enzymes catalyze mineral nitrogen release from complex organics, but also mineral nitrogen, can be produced as the byproduct of organic carbon decomposition. Therefore, a particular nitrogen-mineralization enzyme activity may not always have a negative relation with the available nitrogen (Allison and Vitousek 2005). Above and below a threshold of nutrient availability, negative feedback may be irrelevant to microbial enzyme production. In the case of high nutrient availability, nutrient-mineralization enzyme activity can be low and thus fertilization will not generate additional suppressive effects (Olander and Vitousek 2000). When nutrient availability is low, the nutrient may greatly limit microbial growth and metabolisms. Thus, fertilization may stimulate microbial biomass and in turn microbial enzyme production. Iyyemperumal and Shi (2008) examined N-acetyl-glucosaminidase and protease activities in response to nitrogen fertilization rates. Soil microbial biomass increased with fertilization rates, and enzyme activities were, therefore, greater in fertilized soils than in the unfertilized controls. Substrate induction on enzymes has been attested through substrate addition studies. For example, chitinase activity can be induced by chitin, phosphatase by organic phosphorus, b-glucosidase by cellobiose, and cellulase by cellulose (Smucker and Kim 1987; Chro´st 1991; Shackle et al. 2000). However, positive induction may not always take place. Allison and Vitousek (2005) reported that cellulose addition had no any effect on b-glucosidase activity and also that collagen and cellulose phosphate addition did not increase the activities of respective glycine aminopeptidase and acid phosphatase. Lack of substrate induction on enzymes is thought to occur under some conditions, such as lack of a small amount of available nutrient that is required to activate inducible enzyme synthesis or enzyme

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production at low cost so that enzyme can be produced constantly rather than inducibly (Chro´st 1991; Allison and Vitousek 2005).

3.3.2

Microbial Resource Reallocation

As a cost-efficient entity, the microbial community is expected to allocate resource proficiently for the syntheses of various carbon- and nutrient-acquiring enzymes (Sinsabaugh and Moorhead 1994). In response to a nutrient supply, microbial community likely shifts resource previously used for the synthesis of the nutrientmineralization enzyme to the production of carbon- and other nutrient mineralization enzymes. Some supporting evidence includes increase in phosphatase activity due to improvement in nitrogen availability and increase in cellulose-degrading enzyme activity due to nitrogen and phosphorus fertilization (Dick et al. 1988; Zou et al. 1995; Allison and Vitousek 2005). A resource shift to the production of carbon-mineralization enzymes has been elegantly demonstrated in wetland soils with different available phosphorus and nitrogen (Fig. 3.1, Penton and Newman 2007).

25

High P, Low N

High P, High N

20

Reference Sites Enriched Sites

1.5

log (Ec/Ep)

1.0 0.5 0.0

Low P, Low N

Low P, High N

–0.5 –1.0 –1.5 –1.2

–1.0

–0.8

–0.6

–0.4

–0.2

0.0

0.2

0.4

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log (Ec/En) Fig. 3.1 Log plot of Ec/En vs. Ec/Ep for enriched and reference samples. Each point represents the mean from one sampling period. Ec/Ep values greater than 1 indicate high P availability. Adapted from Penton and Newman (2007)

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In phosphorus and/or nitrogen rich soils, phosphorus- and nitrogen-mineralization enzyme activities were relatively low and instead carbon-mineralization enzyme activity increased. It should be noted that resource partitioning among carbon-, nitrogen-, and phosphorus-mineralization enzymes is an integral feature of microbial community and thus perhaps follows some reaction stoichiometry (Sinsabaugh et al. 2008). Across various ecosystems, for example, some carbon-, nitrogen-, and phosphorus hydrolases are in close 1:1:1 relationship (Sinsabaugh et al. 2008). It is uncertain to what degree microbial resource reallocation for enzyme production can be modulated in terms of relative abundance of organic carbon and nutrient availability. Interestingly, the magnitude of microbial resource allocation appears to be enzyme or nutrient specific. Allison and Vitousek (2005) found that enzymes involved in phosphorus mineralization were more responsive to nitrogen supply than nitrogen-mineralization enzymes to phosphorus availability. Penton and Newman (2007) also observed that carbon-mineralization enzymes were more responsive to phosphorus availability than to nitrogen availability. Effects of nitrogen and phosphorus availability on the production and activity of carbon-mineralization enzymes are intricate. An increase in nitrogen availability often stimulates the activity of cellulase. However, great nitrogen availability is found to reduce the activity of oxidative enzymes such as phenol oxidase and peroxidase.

3.3.3

Microbial Nutrient Demand

Microbial nitrogen mining has been proposed as a microbial process of obtaining nitrogen concealed within recalcitrant organic substances (Moorhead and Sinsabaugh 2006; Craine et al. 2007). Besides in the form of proteins, amino acids, and other labile compounds, nitrogen is tightly associated also with recalcitrant organic carbon. When nitrogen required for microbial growth and metabolisms cannot be met from labile nitrogen mineralization, microbes may “mine” nitrogen from recalcitrant organic matter and plant litters by producing lignin-degrading enzymes. Microbial nitrogen mining hypothesis seems to be in agreement with the common observation that ligninase activity is induced at low nitrogen availability but suppressed at high nitrogen availability (Kirk and Farrell 1987; Fog 1988). For the reason that phosphorous is normally unassociated with organic carbon, microbial phosphorous mining less likely occur (Craine et al. 2007). Microbial nitrogen mining helps to reconcile the suppressive effects of high available nitrogen on lignin-degrading enzymes and predicts that increase in available nitrogen may lead to reduction in decomposition rate.

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W. Shi

Soil Enzyme Activity in Relation to the Quantity and Quality of Soil Organic Matter

As the major carbon and nutrient source, soil organic matter may have positive induction effects on microbial enzyme production. Furthermore, soil enzymes can be immobilized and thus accumulated on soil organic matter. Adsorption appears to provide good protection from decomposition, yet permit the retention of enzyme activity (Burns 1982). Accordingly, soil enzyme activities are expected to be positively related to soil organic matter. Sinsabaugh et al. (2008) examined the relationship between soil organic carbon content and the activity of seven soil enzymes using data from 40 terrestrial ecosystems. They observed that the activities of b-1,4-glucosidase, cellubiohydrolase, b-1,4-N-acetylglucosaminidase, and phosphatase were positively correlated with soil organic carbon content. However, the activity of phenol oxidase, peroxidase, and lucine aminopeptidase was independent of organic carbon content. Soil organic matter is largely heterogeneous in terms of carbon and nutrient chemistry. Provided that substrate positive induction is an important mechanism for soil enzyme production, organic matter chemistry may be manifested in the relative abundance of various carbon and nutrient mineralization enzymes. There are some endeavors for investigating the relationship between organic matter chemistry and the activity of soil enzymes. For example, positive correlations between peptidase and protein and between b-1,4-N-acetyl glucosaminidase and polysaccharides were found in both light and heavy fraction of soil organic carbon (Grandy et al. 2007). In a long-term rye monoculture system, xylanase activity was correlated with the relative abundance of xylan and xylose (Leinweber et al. 2008). Shi et al. (2006) examined the relationship of soil enzyme activities with the chemical composition of soil organic matter in a turfgrass chronosequence. They found that integrated soil enzyme activity was unrelated to chemical compositions of organic matter characterized by Fourier transform infrared spectroscopy. No correlation between enzymes and organic matter is perhaps because soil organic matter reflects longterm decomposition processes whereas enzyme activity fluctuates with current conditions (Grandy et al. 2007).

3.5

3.5.1

Agricultural and Ecological Significance of Soil Enzymes in Soil Carbon Sequestration Assessing Carbon Efflux in Peatlands Under Global Climate Change

Peatlands hold a vast amount of partially decayed vegetation matter due perhaps to phenol oxidase-associated low decomposition rate (Freeman et al. 2001a, 2004).

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Peatland conditions that confine the activity of phenol oxidase, however, are projected to change with global climate change. Because of large organic carbon pool size, small shifts in decomposition rate may exert, through carbon efflux, enormous feedbacks to global environment. Phenol oxidase has been considered as an “enzymatic latch” to hold organic carbon in peatlands (Freeman et al. 2001a). Oxygen that is required for phenol oxidase activity is often absent or limited in peatlands. Upon release of oxygen constraint by frequent droughts, phenol oxidase activity will increase, thereby accelerating the decomposition of soluble phenolics, the compounds inhibitory or toxic to hydrolytic enzymes. In turn, the activity of soil hydrolytic enzymes is improved. Lowering water table in peatlands has been found to significantly increase the activities of hydrolytic enzymes such as b-glucosidase, phosphatase, and sulphatase (Freeman et al. 1996). By eradicating phenol oxidase “latch”, frequent droughts may generate sizeable carbon dioxide efflux into the atmosphere. Furthermore, reduction in organic carbon can impair peatland functions such as food preservation and water purification. Peatlands also contribute greatly to aquatic carbon budget. Approximately, 20% of terrestrial dissolved organic carbon entering the ocean comes from peatlands (Fenner et al. 2007). Global warming can increase dissolved organic carbon export through its controls on phenol oxidase activity (Freeman et al. 2001b; Fenner et al. 2007). An increase of 10
C could lead to 36% increase in the activity of phenol oxidase (i.e., Q10 ¼ 1.36) (Freeman et al. 2001b). In response, dissolved organic carbon increased by a Q10 value of 1.36 and soluble phenolics by a Q10 value of 1.72. Selective enrichment of soluble phenolics may further limit the decomposition of dissolved organic carbon through their inhibitory roles on hydrolytic enzymes, which can strengthen the loss of dissolved organic carbon from peatlands to aquatic systems. Apparently, exports of dissolved organic carbon and soluble phenolics from peatlands are associated with phenol oxidase-degradation effects on peat matrix (Fenner et al. 2005a). Dissolved organic carbon discharge from peatlands can be quite large following a warm and dry period due to great fluctuation in phenol oxidase activity. After a short-term impeded drainage, for example, phenol oxidase activity can be improved and, as a result, soluble phenolics and dissolved organic carbon can increase (Toberman et al. 2008a). Carbon efflux from peatlands to aquatic systems may have significant impacts on regional redistribution of terrestrial carbon and exert significant controls over productivity and biogeochemical cycles in downstream ecosystems (Limpens et al. 2008).

3.5.2

Predicting Soil Carbon Sequestration in Temperate Forests

Lignocellulolytic enzymes have become the focal point to learn about the capacity of soil carbon sequestration in temperate forests under projected atmospheric

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nitrogen deposition. Because of opposite nitrogen effects on cellulose and lignindegrading enzymes, magnitude and direction of nitrogen addition on decomposition appear to depend on the chemistry of organic carbon substances. Carreiro et al. (2000) examined the mass loss of plant litters in relation to relative abundance in lignin and cellulose. For dogwood litters with a low lignocellulose index (i.e., LCI), increase in nitrogen availability from an ambient level to 20 or 80 kg nitrogen ha1 year1 stimulated the activities of both cellulase and phenol oxidase (Fig. 3.2). Cellulase activity alone could explain approximately 65% of variation in litter mass loss. For red oak litters with a high LCI value, increase in nitrogen availability stimulated cellulase activity but repressed phenol oxidase activity. Phenol oxidase alone accounted for about 51% of variation in decay coefficient. Phenol oxidase associated with humified soil organic matter also responds to elevated nitrogen availability. In forests dominated with sugar maples and oaks, phenol oxidase activity in mineral soil was significantly reduced by elevated available nitrogen (Saiya-Cork et al. 2002; DeForest et al. 2004; Gallo et al. 2005), leading to increase in soil organic carbon storage. Over a 3-year period of nitrogen fertilization in temperate forests dominated with sugar maple-red oak and sugar maple-basswood, change in soil organic carbon content was found to be highly and negatively associated with change in soil phenol oxidase activity (Fig. 3.3), indicating the vital role of phenol oxidase in predicting soil carbon sequestration (Waldrop et al. 2004). Waldrop et al. (2004) proposed that ecosystems with highly lignified litters would accrue soil carbon and ecosystems with low lignin content would lose soil carbon in response to increase in soil available nitrogen. Over a three year period, soil carbon decreased by 20% in sugar mapledominant ecosystem and increased by 10% in oak-dominant ecosystem. Although stimulation on cellulolytic enzymes and suppression on oxidative enzymes have been described as the important mechanism dictating the decomposition of plant litters and soil organic matter under elevated nitrogen availability, positive, negative and no effects of nitrogen addition on both oxidative and hydrolytic enzymes have been reported. In general, it is more consistent for nitrogen stimulation effects on the activity of cellulolytic enzymes than for nitrogen suppressive effects on lignin-degrading enzymes (Carreiro et al. 2000; Saiya-Cork et al. 2002; Waldrop et al. 2004; Gallo et al. 2005; Sinsabaugh et al. 2005; Keeler et al. 2009). In addition, magnitude of change in soil enzyme activity in response to nitrogen availability seems to be related to the pool size of accumulative soil enzymes. Given that soil organic matter and clay minerals are two major components in stabilizing extracellular enzymes, enzyme response to an increase in available nitrogen is expected to be less in ecosystems with high soil organic matter and/or clay minerals. Zeglin et al. (2007) demonstrated that increase in nitrogen availability stimulated cellulolytic enzymes by about 40% in a soil with 0.5% of organic carbon, about 10% in a soil with 3% of organic carbon and almost 0% in a soil with 5% organic carbon. Enzyme-based prediction of soil carbon dynamics under atmospheric nitrogen deposition appears to be more significant and detectable in sandy-textured soils (Waldrop et al. 2004). In comparison with finer textured

3 Agricultural and Ecological Significance of Soil Enzymes

a

53

Ambient N +20 kg N.ha –1·yr–1 +80 kg N.ha –1·yr–1

Decay Rate, k (d–1) (X 104)

40

Dogwood

30 20

Maple Oak

10 175 llu 150 la se Ac 125 tiv ity 100

Ce

0.6 0.5 0.4

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b

Decay Rate, k (d–1) (X 104)

40

Dogwood

30 Maple

20

Oak

10

Ph 150 en ol O 100 xid as e Ac t

0.6 0.5 50

0.4

LCI

ivi

ty

Fig. 3.2 Effects of two levels of N addition on the exponential decay rate coefficients (k) of dogwood, red maple, and red oak leaf litters and their relationships with litter LCI (lignocellulose index) and integrated enzyme activities relative to control levels (ambient N): (a) cellulase activity (combined activities of b-glucosidase and cellobiohydrolase); (b) activity of phenol oxidase, a ligninolytic enzyme. Solid bold lines represent the enzyme activities of ambient control levels, defined as 100%. Note especially the switch in direction of the N effect on phenol oxidase activity in the species with the highest LCI (i.e., lignin/[lignin + cellulose]). Adapted from Carreiro et al. (2000)

soil, a sandy soil has a low capacity of adsorption and thus protects soil organic matter poorly from decomposition. Therefore, a large proportion of organic matter can be subjected to enzyme-catalyzed degradation.

W. Shi

Change in soil C (g C/kg soil)

54

1.5 1.0 0.5 0.0 –0.5 –1.0 –1.5 –2.0 –150

y = –0.0057x + 0.0622 R 2 = 0.38 –100

–50

0

50

100

150

200

Change in phenol oxidase activity (µmol·h–1·g–1 OM) Fig. 3.3 Regression analyses of changes in phenol oxidase activity and changes in soil C following high levels of simulated N deposition (80 kg N ha1 year1). OM stands for organic matter. Adapted from Waldrop et al. (2004)

3.5.3

Understanding Soil Carbon Storage Potential in Semi-arid Grasslands

Traditionally, low soil organic matter content in semi-arid grasslands has been attributed to low net primary production and thus low plant carbon input into soil. Until recently, oxidative enzymes have been considered as the culprit for limited accumulation of soil organic matter in semi-arid grasslands (Stursova and Sinsabaugh 2008). The activities of both phenol oxidase and peroxidase are found to be orders of magnitude greater in semi-arid grasslands than in other ecosystems such as forests and grasslands in temperate regions (Stursova et al. 2006; Zeglin et al. 2007). Microbial biomass in a semi-arid grassland in New Mexico was >40% lower than that in temperate grasslands in Kansas, USA and KwaZulu-Natal, South Africa and likewise the activities of soil hydrolytic enzymes including phosphatase, N-acetyl-glucosaminidase, cellobiohydrolase, and b-glucosidase were significantly low (Zeglin et al. 2007). However, the large pool size of oxidative enzymes seems to be unexpected from relatively small microbial biomass. Stursova and Sinsabaugh (2008) proposed that high activity of oxidative enzymes was due to selective stabilization under soil conditions in semi-arid grasslands and as a result, organic matter decomposition could remain at a rate equivalent to the net primary production. Adapting to prolonged dryness and episodic precipitation, ecosystems in a semiarid region may be more susceptible to global environmental change such as rainfall frequency, atmospheric nitrogen deposition, and elevated carbon dioxide. Similar to temperate forests, increase in soil nitrogen availability in semi-arid grasslands can stimulate the activity of soil hydrolytic enzymes such as cellobiohydrolase and b-glucosidase. However, soil nitrogen availability has little impact on the activity of oxidative enzymes (Henry et al. 2005; Stursova et al. 2006; Zeglin et al. 2007). No oxidative enzyme response appears to be a common phenomenon in grasslands where vegetations are of low lignin content (Zeglin et al. 2007). Henry

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et al. (2005) have provided enzyme-based insights on soil carbon dynamics in a semi-arid grassland under global environment change. While atmospheric nitrogen deposition accelerates decomposition via its positive effects on soil hydrolytic enzymes, increase in water availability can accelerate decomposition and thus soil carbon loss via improvement of oxidative enzyme activities.

3.5.4

Restoring Soil Organic Matter from the Perspective of Soil Enzymes

Decline in soil organic matter, which results often from intensive row-crop production, has been recognized as a critical issue limiting long-term land use and thus sustainable agricultural profits. To maintain or improve soil organic matter, diverse management practices such as no till farming, plant residue return, and crop rotations have been increasingly implemented in farming systems. Field-scale trials and observations have provided valuable information on soil organic matter dynamics, under various management practices. However, underlying mechanisms to elucidate management-associated organic matter decomposition appear to be overlooked. Recently, several studies have examined the response of soil lignocellulolytic enzymes to management practices (Henriksen and Breland 1999; Matocha et al. 2004; Zibliske and Bradford 2007). Henriksen and Breland (1999) investigated the effects of soil nitrogen availability on carbon mineralization and soil enzyme activity in wheat straw amended soil in a 70-day incubation experiment. Increase in soil inorganic nitrogen concentration enhanced the activity of cellulolytic enzymes including exocellulase, endocellulase, and hemicellulase and consequently soil carbon mineralization. In wheat straw unamended soil, however, increase in available nitrogen suppressed the activity of endocellulase, hemicellulase, and exocellulase. Such negative effects were thought to result from negative nitrogen effects on humus degradation, thereby reducing the availability of carbohydrates structured within humus complex. In a year-long incubation study, Wang et al. (2004) also found that mineral nitrogen addition stimulated the decomposition during the first 100 days of plant materials of various biochemical compositions, including wheat straw and sugarcane litters. With progressive decomposition, however, negative nitrogen effects on recalcitrant fraction of plant materials appeared. At the end of incubation, mineral nitrogen addition generally had an overall negative effect on decomposition of plant materials. While available nitrogen that is about 1.2% of plant dry matter seems to be sufficient to the optimum decomposition of labile fraction of plan residues (Henriksen and Breland 1999), quantitative information between available nitrogen and its suppressive effect on recalcitrant fraction is limited. Information on phenol oxidase activity in relation to soil nitrogen availability may help to address long-term nitrogen effects on the decomposition of crop residues and soil organic matter. Phenol oxidase appears to be manageable in agricultural soils through application of nitrogen fertilizers. Matocha et al. (2004) reported that nitrogen fertilization

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at 336 kg nitrogen ha1 reduced soil phenol oxidase activity by 38% in no-till systems, but had no effect in the moldboard plow systems. In bermudagrass and tall fescue production systems, activity of soil phenol oxidase was reduced by ammonium nitrate fertilzation at >400 kg nitrogen ha1 year1 and in contrast stimulated by swine lagoon effluent application (Iyyemperumal and Shi 2008). Inconsistent fertilization effects suggest that soil nitrogen availability may not be the sole factor regulating soil phenol oxidase activity. By examining soil phenol oxidase under various oxygen availabilities, Zibliske and Bradford (2007) found that the activity of phenol oxidase could be greatly suppressed under <10% oxygen. In agricultural soil, phenol oxidase appears to act as an “enzymatic latch” to regulate soil organic carbon storage. With the incubation of agricultural soils amended with cowpea or sorghum residues under headspace oxygen concentrations ranging from 0.5 to 21%, the activities of phenol oxidase and peroxidase were found to be negatively related to soluble polyphenolics and dissolved soil organic carbon (Zibliske and Bradford 2007). It appears that reduction in phenol oxidase decreased the decomposition of soluble polyphenolics, which in turn inhibited the decomposition of dissolved organic carbon. These observations agree with those in peatlands and suggest that oxidative enzymes pool can be an important control to the accretion of soil organic carbon in agricultural soils. To develop a management tool from the concept that the activity of oxidative enzymes is an important control for soil carbon storage and nutrient mineralization, more research is needed to quantify the relations of soil oxidative enzymes with soil properties and management practices.

3.6

Conclusions

Soil enzymes have been acknowledged as important soil property to capture soil carbon and nutrient dynamics. Of numerous hydrolytic and oxidative soil enzymes, phenol oxidase appears to be the proximate control dictating the decomposition of soil organic matter. Environment that is conducive to the activity of soil phenol oxidase has been found to favor decomposition of soil organic matter and plant litters and, therefore, likely to impede soil carbon sequestration. To date, several ecosystem perturbation factors and management practices have been evaluated for their impacts on soil phenol oxidase activity. Other soil enzymes that are effective to forecast soil carbon and nutrient dynamics include cellulolytic enzymes and the enzymes for nitrogen and phosphorus mineralization. As a cost-efficient entity, microbial community appears to possess several tactics for producing extracellular enzymes in relation to organic carbon chemistry and nutrient availability. End product suppression on enzymes represents one of microbial strategies and accounts for negative associations observed quite often between the activity of soil phosphatases and phosphorus availability and in some cases between the activity of N-acquiring enzymes and nitrogen availability. Nutrient induction for carbon- and other nutrient-mineralization enzymes reflects microbial

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resource reallocation in response to a nutrient supply. This strategy predicts that the activities of carbon- and phosphorous-mineralization enzymes will increase with nitrogen fertilization and the activities of carbon- and nitrogen-mineralization enzymes can be strengthened under the condition of abundant phosphate. Microbial nitrogen mining emerges from the observation that phenol oxidase activity is inhibited by an increase in available nitrogen. An ability of mining a nutrient concealed within recalcitrant organic substances seems to be another approach for microbial community to generate some carbon-mineralization enzymes. There are a number of robust cases that soil enzyme activities can be predicted from microbial strategies for extracellular enzyme production and that changes in soil enzyme activities are able to manifest the direction and magnitude of organic carbon decomposition and nutrient mineralization. In forest ecosystems, for example, increase in cellulase activity and decrease in phenol oxidase activity under projected atmospheric nitrogen deposition seem to be in agreement with predictions based on microbial resource allocation and microbial nitrogen mining. Modifications in lignocellulolytic enzyme activities are, therefore, accountable for changes in decomposition rates and quantity of soil carbon retention. In peatlands, changes in phenol oxidase activity are tightly associated with carbon efflux to atmosphere and hydrosphere. However, there are some instances that neither microbial strategies for enzyme production nor relationships of soil enzyme activities with decomposition and mineralization are as expected. In order to reconcile different observations, some uncertainties need to be addressed, including thresholds for nutrient availability that determines whether nutrient suppression or induction on enzymes can take place, the relative abundance of soil organic matter and/or clay minerals that sustain a sufficient pool size of soil enzymes and therefore mask any effects associated with slight pool size change, and organic matter chemistry that determine the relative importance of cellulolytic enzymes and phenolics-degrading enzymes for predicting soil carbon decomposition. Mechanisms that underline microbial enzyme production and the predictability of soil enzymes for soil carbon and nutrient dynamics have great implications in terms of global environmental change as well as management practices. To restore soil organic matter and optimize nutrient cycling in agricultural systems, we need to develop management practices that take the principles of soil enzyme activities into considerations.

References Alef K, Nannipieri P (1995) Cellulase activity. In: Alef K, Nannipieri P (eds) Methods in applied soil microbiology and biochemistry. Academic, San Diego, CA, pp 345–349 Alef K, Nannipieri P, Trazar-Cepeda C (1995) Phosphatase activity. In: Alef K, Nannipieri P (eds) Methods in applied soil microbiology and biochemistry. Academic, San Diego, CA, pp 335–344 Allison SD, Vitousek PM (2005) Responses of extracellular enzymes to simple and complex nutrient inputs. Soil Biol Biochem 37:937–944

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Allison SD, Czimczik CI, Treseder KK (2008) Microbial activity and soil respiration under nitrogen addition in Alaskan boreal forest. Global Change Biol 14:1156–1168 Bandick AK, Dick RP (1999) Filed management effects on soil enzyme activities. Soil Biol Biochem 31:1471–1479 Burke M, Cairney JWG (2002) Laccases and other polyphenol oxidases in ecto- and ericoid mycorrhizal fungi. Mycorrhiza 12:105–116 Burns RG (1982) Enzyme activity in soil: location and a possible role in microbial ecology. Soil Biol Biochem 14:423–427 Carreiro MM, Sinsabaugh RL, Repert DA, Parkhurst DF (2000) Microbial enzyme shifts explain litter decay responses to simulated nitrogen deposition. Ecology 81:2359–2365 Chro´st RJ (1991) Environmental control of the synthesis and activity of aquatic microbial ectoenzymes. In: Chro´st RJ (ed) Microbial enzymes in aquatic environments. Springer, New York, pp 29–59 Claus H (2004) Laccases: structure, reactions, distribution. Micron 35:93–96 Craine JM, Morrow C, Fierer N (2007) Microbial nitrogen limitation increases decomposition. Ecology 88:2105–2113 Criquet S, Tagger S, Vogt G, Iacazio G, Le Petit J (1999) Laccase activity of forest litter. Soil Biol Biochem 31:1239–1244 DeForest JL, Zak DR, Pregitzer KS, Burton AJ (2004) Atmospheric nitrate deposition and the microbial degradation of cellobiose and vanillin in a northern hardwood forest. Soil Biol Biochem 36:965–971 Dick RP (1992) A review: long-term effects of agricultural systems on soil biochemical and microbial parameters. Agr Ecosyst Environ 40:25–36 Dick RP, Rasmussen PE, Kerle EA (1988) Influence of long-term residue management on soil enzyme activities in relation to soil chemical properties of a wheat-fallow system. Biol Fertil Soils 6:159–164 Dodor DE, Tabatai MA (2007) Arylamidase activity as an index of nitrogen mineralization in soils. Commun Soil Sci Plan 38:2197–2207 Ekenler M, Tabatabai MA (2002) Beta-glucosaminidase activity of soils: effect of cropping systems and its relationship to nitrogen mineralization. Biol Fertil Soils 36:367–376 Ekenler M, Tabatabai MA (2004) Beta-glucosaminidase activity as an index of nitrogen mineralization in soils. Commun Soil Sci Plan 35:1081–1094 Endo K, Hayaski Y, Hibi T, Hosono K, Beppu T, Ueda K (2003) Enzymological characterization of EpoA, a laccase-like phenol oxidase produced by Streptomyces griseus. J Biochem 133:671–677 Fenner N, Freeman C, Reynold B (2005a) Observations of a seasonally shifting thermal optimum in peatland carbon-cycling processes: implications for the global carbon cycle and soil enzyme methodologies. Soil Biol Biochem 37:1814–1821 Fenner N, Freeman C, Reynolds B (2005b) Hydrological effects on the diversity of phenolic degrading bacteria in a peatland: implications for carbon cycling. Soil Biol Biochem 37:1277–1287 Fenner N, Freeman C, Lock MA, Harmens H, Reynolds B, Sparks T (2007) Interactions between elevated CO2 and warming could amplify DOC exports from peatland catchments. Environ Sci Technol 41:3146–3152 Finzi AC, Sinsabaugh RL, Long TM, Osgood MP (2006) Microbial community responses to atmospheric carbon dioxide enrichment in a warm-temperate forest. Ecosystems 9:215–226 Fog K (1988) The effect of added nitrogen on the rate of decomposition of organic matter. Biol Rev 63:433–462 Freeman C, Liska G, Ostle NJ, Lock MA, Reynolds B, Hudson J (1996) Microbial activity and enzymic decomposition processes following peatland water table drawdown. Plant Soil 180:121–127 Freeman C, Ostle N, Kang H (2001a) An enzymic ‘latch’ on a global carbon storage. Nature 409:149

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Freeman C, Evans CD, Monteith DT, Reynolds B, Fenner N (2001b) Export of organic carbon from peat soils. Nature 412:785 Freeman C, Olstle NJ, Fenner N, Kang H (2004) A regulatory role for phenol oxidase during decomposition in peatlands. Soil Biol Biochem 36:1663–1667 Gallo ME, Lauber CL, Cabaniss SE, Waldrop MP, Sinsabaugh RL, Zak DR (2005) Soil organic matter and litter chemistry response to experimental N deposition in northern temperate deciduous forest ecosystems. Global Change Biol 11:1514–1521 Grandy AS, Neff JC, Weintrau MN (2007) Carbon structure and enzyme activities in alpine and forest ecosystems. Soil Biol Biochem 39:2701–2711 Hammel KE (1997) Fungal degradation of lignin. In: Cadish G, Giller KE (eds) Driven by nature. CAB International, London, pp 33–45 Henriksen TM, Breland TA (1999) Nitrogen availability effects on carbon mineralization, fungal and bacterial growth, and enzyme activities during decomposition of wheat straw in soil. Soil Biol Biochem 31:1121–1134 Henry HA, Juarez JD, Field CB, Vitousek PM (2005) Interactive effects of elevated CO2, N deposition and climate change on extracellular enzyme activity and soil density fractionation in a California annual grassland. Global Change Biol 11:1808–1815 Hullo MF, Moszer I, Danchin A, Martin-Verstraete I (2001) CoA of Bacillus subtilis is a copperdependent laccase. J Bacteriol 183:5426–5430 Iyyemperumal K, Shi W (2008) Soil enzyme activities in two forage systems following application of different rates of swine lagoon effluent or ammonium nitrate. Appl Soil Ecol 38:128–136 Kang H, Freeman C (1999) Phosphatase and arylsulphatase activities in wetland soils: annual variation and controlling factors. Soil Biol Biochem 31:449–454 Keeler BL, Hobbie SE, Kellogg LE (2009) Effects of long-term nitrogen addition on microbial enzyme activity in eight forested and grassland sites: implications for litter and soil organic matter decomposition. Ecosystems 12:1–15 Kirk TK, Farrell RL (1987) Enzymatic combustion: the microbial degradation of lignin. Annu Rev Microbiol 41:465–505 Ladd JN (1978) Origin and range of enzymes in soil. In: Burns RG (ed) Soil enzymes. Academic, New York, pp 51–96 Leinweber P, Jandl G, Baum C, Eckhardt KU, Kandeler E (2008) Stability and composition of soil organic matter control respiration and soil enzyme activities. Soil Biol Biochem 40:1496–1505 Limpens J, Berendse F, Blodau C, Canadell JG, Freeman C, Holden J, Noulet N, Rydin H, Schaepman-Strub G (2008) Peatlands and the carbon cycle: from local processes to global implications – a synthesis. Biogeosciences 5:1475–1491 Lipson DA, Wilson RF, Oechel WC (2005) Effects of elevated atmospheric CO2 on soil microbial biomass, activity and diversity in a charparral ecosystem. Appl Environ Microb 71:8573–8580 Matocha CJ, Haszler GR, Grove JH (2004) Nitrogen fertilization suppresses soil phenol oxidase enzyme activity in no-tillage systems. Soil Sci 169:708–714 McLatchey GP, Reddy KR (1998) Regulation of organic matter decomposition and nutrient release in a wetland soil. J Environ Qual 27:1268–1274 Moorhead DL, Sinsabaugh RL (2006) A theoretical model of litter decay and microbial interaction. Ecol Monogr 76:151–174 Olander LP, Vitousek PM (2000) Regulation of soil phosphatase and chitinase activity by N and P availability. Biogeochemistry 49:175–190 Paul EA, Clark FE (1996) Soil microbiology and biochemistry. Academic, San Diego, CA Penton CR, Newman S (2007) Enzyme activity responses to nutrient loading in subtropical wetlands. Biogeochemistry 84:83–98 Pind A, Freeman C, Lock MA (1994) Enzymatic degradation of phenolic materials in peatlands – measurement of phenol oxidase activity. Plant Soil 159:227–231 Saiya-Cork KR, Sinsabaugh RL, Zak DR (2002) The effects of long-term nitrogen deposition on extracellular enzyme activity in an Acer saccharum forest soil. Soil Biol Biochem 34:1309–1315

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Schimel JP, Weintraub MN (2003) The implications of exoenzyme activity on microbial carbon and nitrogen limitation in soil: a theoretical model. Soil Biol Biochem 35:549–563 Shackle VJ, Freeman C, Reynolds B (2000) Carbon supply and the regulation of enzyme activity in constructed wetlands. Soil Biol Biochem 32:1935–1940 Shi W, Dell E, Bowman D, Iyyemperumal K (2006) Soil enzyme activities and organic matter composition in a turfgrass chronosequence. Plant Soil 288:285–296 Sinsabaugh RL (1994) Enzymic analysis of microbial pattern and process. Biol Fertil Soils 187:69–74 Sinsabaugh RL, Linkins AE (1993) Statistical modeling of litter decomposition from integrated cellulase activity. Ecology 74:1594–1597 Sinsabaugh RL, Moorhead DL (1994) Resource allocation to extracellular enzyme production: a model for nitrogen and phosphorous control of litter decomposition. Soil Biol Biochem 26:1305–1311 Sinsabaugh RL, Antibus RK, Linkins AE, McClaugherty CA, Rayburn L, Repert D, Weiland T (1992) Wood decomposition over a 1st-order watershed – mass-loss as a function of lignocellulase activity. Soil Biol Biochem 24:743–749 Sinsabaugh RL, Antibus RK, Linkins AE, MaClaugherty CA, Rayburn L, Repert D, Weiland T (1993) Wood decomposition – nitrogen and phosphorus dynamics in relation to extracellular enzyme activity. Ecology 74:1586–1593 Sinsabaugh RL, Gallo ME, Lauber C, Waldrop MP, Zak DR (2005) Extracellular enzyme activities and soil organic matter dynamics for northern hardwood forests receiving simulated nitrogen deposition. Biogeochemistry 75:201–215 Sinsabaugh RL, Lauber CL, Weintraub MN, Ahmed B, Allison SD, Crenshaw C, Contosta AR, Causack D, Frey S, Gallo ME, Gartner TB, Hobbie SE, Holland K, Keeler BL, Powers JS, Stursova M, Takacs-Vesbach C, Wallenstein MD, Zak DR, Zeglin LH (2008) Stoichiometry of soil enzyme activity at global scale. Ecol Lett 11:1252–1264 Smucker RA, Kim CK (1987) Chitinase induction in an estuarine ecosystems. In: Llewellyn GC, O’Rear CE (eds) Biodegradation research. Plenum, New York, pp 347–355 Speir TW, Ross DJ (1978) Soil phosphatase and sulphatase. In: Burns RG (ed) Soil enzymes. Academic, New York, pp 197–250 Stursova M, Sinsabaugh RL (2008) Stabilization of oxidative enzymes in desert soil may limit organic matter accumulation. Soil Biol Biochem 40:550–553 Stursova M, Crenshaw CL, Sinsabaugh RL (2006) Microbial responses to long-term N deposition in a semiarid grassland. Microb Ecol 51:90–98 Tabatabai MA (1994) Soil enzymes. In: Weaver RW, Angle S, Bottomley P, Bezdicek D, Smith S, Tabatabai A, Wollum A (eds) Methods of soil analysis, part 2 – microbiological and biochemical properties. Soil Science Society of America Inc, Wisconsin, pp 775–833 Toberman H, Artz FC, RRE ECD, Fenner N (2008a) Impeded drainage stimulates extracellular phenol oxidase activity in riparian peat cores. Soil Use Manage 24:357–365 Toberman H, Evans CD, Freeman C, Fenner N, White M, Emmett BA, Artz RRE (2008b) Summer drought effects upon soil and litter extracellular phenol oxidase activity and soluble carbon release in an upland Calluna heathland. Soil Biol Biochem 40:1519–1532 Waldrop MP, Zak DR, Sinsabaugh RL, Gallo M, Lauber C (2004) Nitrogen deposition modifies soil carbon storage through changes in microbial enzymatic activity. Ecol Appl 14:1172–1177 Wang WJ, Baldock JA, Dalal RC, Moody PW (2004) Decomposition dynamics of plant materials in relation to nitrogen availability and biochemistry determined by NMR and wet-chemical analysis. Soil Biol Biochem 36:2045–2058 Zeglin LH, Stursova M, Sinsabaugh RL, Collins SL (2007) Microbial responses to nitrogen addition in three contrasting grassland ecosystems. Oecologia 154:349–359 Zibliske LM, Bradford JM (2007) Oxygen effects on carbon, polyphenols, and nitrogen mineralization potential in soil. Soil Sci Soc Am J 71:133–139 Zou X, Binkley D, Caldwell BA (1995) Effects of dinitrogen-fixing trees on phosphorus biogeochemical cycling in contrasting forests. Soil Sci Soc Am J 59:1452–1458

Chapter 4

Enzymes in Forest Soils Petr Baldrian and Martina Sˇtursova´

4.1

Introduction

Forest soils typically accumulate dead organic material, known as plant litter, on the soil surface. When fresh, this material contains a variety of substrates, such as soluble saccharides, organic acids, amino acids or starch, as well as the plant cell wall-derived biopolymers, cellulose, hemicelluloses and lignin, which are utilized during the growth of soil decomposer microorganisms. The sequential transformation of plant litter in temperate forests leads to the formation of a distinct organic horizon derived from the transformation products of litter decomposition as well as the dead biomass of soil microorganisms. The organic horizon is known for its high content of humic substances. Compared to agricultural soils and grassland soils, forest soils contain more fungal biomass (van der Wal et al. 2006), and the mycorrhizal community is formed mainly by the ectomycorrhizal fungi associated with tree roots instead of the arbuscular mycorrhizal fungi, which dominate the grasslands. Due to the high input of organic matter, enzymes participating in the decomposition of organic matter play an important role in forest soils. The enzymes relevant to decomposition studies are those related to the degradation of plant cell wall polymers (cellulose, hemicellulose, lignin, pectin) and microbial biomass (chitin and other polysaccharides). Besides these, enzyme systems involved in the acquisition of nitrogen, phosphorus, and sulfur are also of interest because these elements are indispensable for the growth of soil microbial biomass. Plant litter decomposition is actually the most important process in shaping the formation of the soil matter gradient. This complex process involves numerous taxa of decomposer microorganisms and a wide array of enzymatic transformations of

P. Baldrian (*) and M. Sˇtursova´ Laboratory of Environmental Microbiology, Institute of Microbiology of the ASCR, v.v.i, Vı´denˇska´ 1083, Praha 4 14220, Czech Republic e-mail: [email protected]

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individual molecules. It has been described as a successional process, where the initial chemical composition defines the microbial community which, in turn, secretes degradative enzymes. The produced enzymes change the chemical properties of the substrate. These changes further drive the decomposer community succession in the litter (Sinsabaugh et al. 2002a). In this chapter, factors affecting the activity of enzymes in forest soils with respect to their spatial distribution are discussed. The lignocellulose-degrading enzymes and their producers are the subject of another chapter in this book (Chap. 9).

4.2

Distribution of Enzymes in the Soil Profile

The stratification of organic compounds due to their sequential transformation by decomposer microorganisms leads to the formation of a sharp gradient of soil properties in the upper layers of forest soils. Along with the amount of total soil carbon and its readily utilizable forms, microbial biomass and activity (e.g., respiration) typically decrease with soil depth (Sˇnajdr et al. 2008b). This results in a sharp vertical gradient of activities of lignocellulose-degrading enzymes as well as of other enzymes (chitinases, phosphatases, peptidases, proteases, catalase, and invertase). This type of vertical distribution of enzyme activities was reported in several forest soils ranging from the Mediterranean evergreen forests and temperate forests to boreal ecosystems (Trasar-Cepeda et al. 2000; Prietzel 2001; Andersson et al. 2004; Wittmann et al. 2004; Zhang et al. 2004). Recent results on the vertical distribution of enzymes in a temperate Quercus petraea forest demonstrated that enzyme activities dramatically change with soil depth, even within individual soil horizons. Most apparently, the upper part of the organic horizon is significantly different from the lower part with respect to the organic carbon content, enzyme activities and microbial biomass (Sˇnajdr et al. 2008b; Fig. 4.1). Although the spatial differences of enzyme activities level-off in the deeper parts of soils, the activity of phosphatase, arylsulfatase, protease, and urease decrease with soil depth and soil organic matter down to depths greater than 1 m (Venkatesan and Senthurpandian 2006). Microbial biomass analyses showed that soil is also stratified with respect to the relative abundance of individual microbial groups. The relative amount of fungal biomass tends to decrease with soil depth, while the relative amount of actinomycetes increases (Baldrian et al. 2008; Sˇnajdr et al. 2008b). This is reflected in the differences in the relative abundance of individual enzymes at different depths, since these partly reflect the abundance of their producers. Some studies have shown that enzymes targeting easily decomposable compounds (e.g., invertase or proteases) can be limited to the surface soil layers, while oxidases and peroxidases are significantly active at great depths (Zhang et al. 2005; Venkatesan and Senthurpandian 2006). While most enzymes show higher activity in the topsoil horizons, the highest arylsulfatase activity in a boreal pine forest was recorded in the deeper horizons where bacterial biomass dominated over fungal biomass (Wittmann et al. 2004).

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Fig. 4.1 The vertical gradient of soil organic matter content, microbial biomass, respiration and the activity of selected enzymes in a hardwood forest soil profile (Quercus petraea, Czech Republic, Europe) shows that enzyme activity decreases in the soil profile along with a decrease in soil organic matter content, soil microbial biomass and microbial activity. Bacterial biomass is based on the amount of bacteria-specific PLFA, and fungal biomass is based on ergosterol content. Depth was measured below the litter horizon. Abbreviations: Lac laccase, MnP Mn-peroxidase, EG Endo-1,4-b-glucanase, CBH cellobiohydrolase, bG b-glucosidase, Chit N-acetylglucosaminidase, AcP acid phosphatase. Data from Sˇnajdr et al. (2008b)

Litter quality and quantity is primarily responsible for the formation of soil property gradients with different slopes. In the same soil, the decrease of soil microbial biomass content and enzyme activity with depth was more pronounced under a hardwood tree (Robinia pseudoacacia) than in the same soil with a softwood Pinus sylvestris, the litter from which is more difficult to decompose (Landgraf et al. 2005).

4.3

Spatial Variability of Enzyme Activities in Soils

In contrast to uniform agricultural soils that are homogenized during land management, forest soils exhibit a significant level of horizontal variability in their properties. Relatively low variability is usually observed for physico-chemical variables, with coefficients of variation (CVs) of 5–10% for pH and approximately 25% for organic carbon, total carbon and total nitrogen in Quercus spp. and Fagus spp. soils (Morris 1999; G€om€ oryova´ 2004). Microbial biomass content is usually more variable: A CV of 44% was reported for fungal biomass content in a Pinus sylvestris forest humus layer and CVs of 36–48% were reported for both fungal and bacterial biomass in the upper 5 cm of a Quercus rubra forest soil (Morris 1999; M€ott€onen et al. 1999); CVs for individual PLFA molecules in the upper 5 cm of Picea abies/

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Betula pubescens soil ranged from 16 to 37% (Saetre and Baath 2000). Soil enzyme activities are more widely varied and the recorded values often span several orders of magnitude (Prietzel 2001; G€ om€ oryova´ 2004; Luis et al. 2005; Sˇnajdr et al. 2008b). For enzyme activities in Quercus spp. forest soils, the CVs were similar in the H and Ah horizons and ranged from 29 to 64% for phosphatase, b-glucosidase, endoglucanase, and other enzymes (Trasar-Cepeda et al. 2000; Sˇnajdr et al. 2008b). The two most apparent factors that may affect the distribution of enzyme activities in soil are (1) the presence of their microbial producers and (2) the presence of substrates or inducers of enzyme expression. Enzymes produced constitutively by all groups of microorganisms, such as acid phosphatase (Criquet et al. 2004), can thus be expected to correlate with soil microbial biomass. Microbial carbon, respiration, and chitinase were shown to correlate in the H horizon of spruce and beech forest soil, but cellulase was not (Andersson et al. 2004). In studies with Quercus ilex litter, bacterial and fungal counts correlated with the activities of selected glycosyl hydrolases, but not with ligninolytic enzymes (Criquet et al. 2000, 2002). Fungi are often considered to be responsible for the production of several oxidative and hydrolytic enzymes in soils and the variation of fungal abundance is thus one of the important factors shaping the distribution of enzymes in soils (Moller et al. 1999; Caldwell 2005; Baldrian 2008). It was demonstrated in soil microcosms that the presence of saprotrophic basidiomycetes increased the activities of ligninolytic enzymes, polysaccharide hydrolases, chitinase, and acid phosphatase compared to non-colonized soil (Sˇnajdr et al. 2008a). Elevated activities of several enzymes were also recorded in soils under mycelia of fungi (Gramss 1997). Soil quality is the second major factor that can potentially affect enzyme activity. In a mixed forest, bacterial biomass and community composition are regulated by the proximity of different tree species, presumably caused by the differences in litter quality, rhizosphere effects or the variation of soil moisture due to differential throughfall (Saetre 1999; Saetre and Baath 2000). In a Fagus sylvatica forest, catalase activity exhibited a patchy distribution with most of the activity found at tree crown perimeters (G€ om€ oryova´ 2004). In contrast, phosphatase activity in rhizosphere soil of fir forests was found to be higher near tree stems as compared to the soil further from stems, which is probably due to the higher phosphate demand of tree root-colonized soil compared to bulk soil (Chen 2003). In an attempt to locally detect enzyme activities in soils using immobilized substrates, it was demonstrated that the activity of several hydrolytic enzymes (e.g., phosphatase, aminopeptidase, chitinase, or b-glucosidase) in coniferous forest soils frequently co-localize with living biomass in the soil profile, such as roots, mycelial mats or ectomycorrhizal systems (Dong et al. 2007). In ectomycorrhizas, the expression of enzymes differs between the hartig net, fungal cords and fine mycelium. High phosphatase activity of fungal origin was found in fine Suillus hyphae and in mycorrhizal roots (Timonen and Sen 1998). Cellulase and chitinase decreased with the decreasing C/N ratio in spruce litter and humus, but increased at the N-rich sites of beech litter microcosms (Andersson et al. 2004). An explanation for this may be that the decrease in the C/N ratio in litter corresponds with a decrease in accessible carbon sources in the

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soil during litter transformation, while extra nitrogen added to the litter increases its degradation. Humus under Alnus spp. with high nitrogen content exhibited higher microbial biomass and higher enzyme activities than humus under pine growing in the same type of soil (Niemi et al. 2007).

4.4 4.4.1

Factors Affecting Soil Enzyme Activities Producers of Extracellular Enzymes

As mentioned above, fungi seem to be the only group of microorganisms that produce significant amounts of ligninolytic enzymes in soils and also seem to be the major producers of polysaccharide hydrolases (Moller et al. 1999; Baldrian 2008). This fact is underlined in forest soils by the high proportion of fungal biomass compared to other ecosystems. It was also demonstrated that fungi are responsible for the production of chitinase (N-acetylglucosaminidase) in soils, while the activity of arylsulfatase seems to be more reflective of the distribution of bacterial biomass (Moller et al. 1999; Baldrian et al. 2008). However, some enzymes, like phosphatase or b-glucosidase, are produced by a wide spectrum of soil organisms and their activity mainly corresponds to the amount of soil organic matter and microbial biomass (Amador et al. 1997).

4.4.2

Soil Chemistry and Nutrient Availability

The addition of low molecular mass organic compounds, such as monosaccharides or organic acids, affects the activity of several enzymes, such as acid phosphatase, phosphodiesterase, and urease, while other enzymes, such as protease, are unaffected. These additions also increase respiration (Renella et al. 2007). b-Glucosidase and phosphatase were also increased by the addition of carbon or nitrogen (Allison and Vitousek 2005). Although the available phosphorus content in natural soil does not affect phosphatase activity (Venkatesan and Senthurpandian 2006), the lower demand for phosphorus after its addition typically leads to a decrease in phosphatase activity (Olander and Vitousek 2000). Similar results were also found for arylsulfatase in hardwood forest soil where the enzyme negatively correlated with SO42 concentration and decreased by tens of percent after ammonium sulfate addition (Prietzel 2001). The addition of cellulose increases both cellulolysis and total mineralization in soils, probably by the activation of cellulolytic enzymes. Although xylanase is often co-localized with cellulases in soils (Sˇnajdr et al. 2008b), it is not induced by the addition of cellulose (Scheu et al. 1993).

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The effect of adding nitrogen to soils was frequently studied due to the fact that this process accompanies the anthropogenic pollution of the environment. Studies on lignocellulose degradation after nitrogen supplementation indicated that the effect depends greatly on the properties of the ecosystem. For more information, please see Chap. 9 of this book. Chitinase activity was found to negatively correlate with soil nitrogen content and nitrogen addition decreased chitinase activity in soil with low nitrogen content (Olander and Vitousek 2000; Allison et al. 2008). This effect is probably a result of the fact that nitrogen addition decreases the relative amount of fungi in soils. There are, however, studies where neither fungal biomass content nor chitinase activity responded to simulated nitrate addition (Waldrop and Zak 2006) and the effect seems to be dependent on the properties of the ecosystem. In general, the addition of nitrogen to hardwood forest soil has a more pronounced effect in the litter horizon than in the organic horizon. Urease, phosphatase, and several glycosidases increased by 7–56% in N-supplemented plots, while laccase and chitinase responded differently in soil and litter (Saiya-Cork et al. 2002). The effect of elevated nitrogen input into soils is also apparent in cases where forests are invaded by trees with nitrogen-fixing symbionts, such as Alnus spp. or Falcataria spp. In both cases, a shift in the soil microbial community composition leading to the dominance of bacteria over fungi in high nitrogen conditions accompanied the changes in enzyme activities (Selmants et al. 2005; Allison et al. 2006). The succession of vegetation and microbial communities is an important process, occurring in both natural and cultured ecosystems. The chemical changes accompanying the formation of a soil horizon are also reflected by the changes in microbial biomass and enzyme activity in top soils. Activities of extracellular enzymes and microbial biomass in topsoil tended to increase for at least 20 years during the spontaneous succession of coal mine deposits in temperate zones (Baldrian et al. 2008). In managed spruce forests, invertase and peroxidase activities increased until the site reached 23 years of age (Zhang et al. 2004). In both cases, the enzyme activities and microbial biomass decreased in the late stages of succession. In tropical soils that were reclaimed after bauxite mining, microbial biomass and enzyme activities increased much more rapidly after recovery and reached a plateau value after only 1 year of treatment; the increase of carbon and nitrogen content in the same soils was considerably slower and lasted for several years (Carneiro et al. 2008).

4.4.3

Seasonality

In biomes with substantial variation in temperature or moisture during the year, activities of extracellular enzymes in forest soils and litter show considerable seasonal variation. In previous studies, climatic factor such as temperature and soil moisture content have been identified as the main responsible factor for the observed seasonal differences in the case of laccase, polysaccharide hydrolases,

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phosphatase, urease, protease, and other enzymes (Dilly and Munch 1996; Criquet et al. 2000, 2002, 2004; Wittmann et al. 2004; Baldrian et al. 2008; Bastida et al. 2008). In hardwood forest with the litterfall season limited to autumn, the changes in the chemical composition of litter during its decomposition are another important cause of seasonal differences in enzyme activities (Dilly and Munch 1996; Fioretto et al. 2000). In soils of hardwood forests developed during succession on mine deposits, seasonality was actually the most important factor in determining the variability of enzyme activities. Seasonality had a significant effect on glycosidases, arylsulfatase, chitinase, and especially phosphatase, accounting for 10–70% of the observed variability (Baldrian et al. 2008). In temperate zones, higher enzyme activities are usually detected in the warm summer period as compared to the winter (Baum and Hrynkiewicz 2006; Niemi et al. 2007). Although the winter period is often regarded as a time of limited microbial activity in boreal forests where there are long periods of low soil temperature, as much as 7–32% of the enzyme-catalyzed annual turnover of organic compounds was estimated to occur during the 6 cold months of the year (Kahkonen et al. 2001; Wittmann et al. 2004). In ectomycorrhizal fungi, the activity of several hydrolases (particularly phosphatase and b-glucosidase) was higher in the winter when the supply of photosynthates from the host tree was limited as compared to the spring, resulting in a seasonal response of enzyme production based on nutrient availability (Mosca et al. 2007).

4.4.4

Soil Moisture Content

Soil moisture was previously reported to affect the microbial biomass, respiration in the soil and enzyme-catalyzed processes (Orchard and Cook 1983; Criquet et al. 2002). Most of the previous studies, however, focused on the effects of temporary drying and not on the effect of spatial variability. In dry periods, forest soils were reported to contain less microbial biomass and exhibited reduced enzyme activities (Ross 1987; Sardans and Penuelas 2005). In Mediterranean forests, the activities of laccase, b-glucosidase, acid phosphatase, urease, and protease were substantially reduced in the dry seasons (Criquet et al. 2002, 2004; Sardans and Penuelas 2005), and Mn-peroxidase in evergreen oak litter was only detected during humid periods (Criquet et al. 2000). There is, however, also a significant level of spatial variation in soil moisture at any given time. The coefficients of variation of moisture in the upper part of different temperate forest soils were reported to be 10–60%, and the range of moisture content varied between 15 and 70%. The spatial differences in soil moisture content can partly to ascribed to the differences in throughfall (and the tree cover, since throughfall strongly correlates with the gap fraction) partly to the consumption by plant roots or translocation by fungal mycelia and partly to simple physical draining.

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The spatial variation of soil moisture was found to affect the total microbial biomass (Stoyan et al. 2000; G€ om€ oryova´ et al. 2006), although not necessarily the entire microbial community; in Quercus rubra forest soil, only bacterial and not fungal biomass was affected (Morris 1999). The spatial variability of soil moisture content also affects microbial processes. In different forest types, the moisture content positively correlated with the rate of N-transformation (Bengtson et al. 2005), phosphatase or catalase activity (Kramer and Green 2000; G€om€oryova´ et al. 2006). The effect of drying is very important since a relatively small moisture content reduction of 10% in Mediterranean evergreen forests caused 10–80% of the urease, protease and b-glucosidase activity. A higher reduction (21%) in moisture content caused by rainfall and runoff exclusion had an even more pronounced effect on the enzymes (Sardans and Penuelas 2005).

4.5

Litter Decomposition in Forest Ecosystems

Litter decomposition is the fastest process of soil organic matter decomposition due to the high initial amount of soluble and polymeric polysaccharides in the senescent plant material. Fresh leaf litter is typically composed of 3–7% soluble polysaccharides, 15–27% cellulose and 15–50% lignin. During litter transformation, soluble compounds including polysaccharides are removed first and lignin, which is degraded more slowly, usually accumulates (Osono and Takeda 2005). These changes in litter quality are reflected in the sequential production of individual enzymes participating in the decomposition of different litter components. Enzymes that act on soluble saccharides like invertase or a-glucosidase and b-glucosidase activity usually peak in the initial stages of decomposition (Sinsabaugh et al. 2002a; Fioretto et al. 2007; Papa et al. 2008). The activities of other cellulases usually increase more slowly and peak when the mass loss is at 40–80%. The activity of phenol oxidizing enzymes typically peak at late stages of decomposition when the available cellulose disappears (Sinsabaugh et al. 2002a). The higher laccase activity recorded during the late stages of decomposition on Quercus ilex and Myrtus communis litter was associated with an increase in the fungal biomass content of the litter (Fioretto et al. 2007; Papa et al. 2008). The successive production of enzymes on decomposing litter reflects the successional changes of the decomposer community. With respect to fungi, it is generally accepted that the proportion of basidiomycetes that are able to produce ligninolytic enzymes increases with time, while ascomycetes dominate the earlier stages of decomposition, with many of them already being present in the senescent litter as endophytes (Aneja et al. 2006; Osono 2007). In beech litter degradation experiments, fungi colonizing fresh litter were mainly pectinase producers, while the middle phases of decomposition were dominated by cellulase-producing species and the late phases were characterized by cellulolytic and chitinolytic fungi (Kjoller and Struwe 1990, 2002).

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Water soluble C content, cellobiohydrolase and laccase activities were found to correlate with mass loss in different litter types (Allison and Vitousek 2004). Linkins et al. (1990a, b) followed the decomposition of Cornus florida, Quercus prinus, and Acer rubrum leaves in a deciduous woodland and in vitro. Enzyme activities were affected by litter type and both the total mass loss and the loss of cellulose correlated with cellulase activities. However, clearly linking enzyme activity to litter decomposition is not always so simple. The decomposition of Cornus florida litter was accomplished with much less enzyme activity than that needed for the same mass loss in Quercus prinus or Acer rubrum litter (Carreiro et al. 2000). Litter decomposition is a sensitive process which is affected by several external factors including forest management, nutrient supply and the presence and activity of microinvertebrates (Schlatte et al. 1998; Sinsabaugh et al. 2002b; Waldrop et al. 2003). Litter decomposition is also subject to seasonality. During the decomposition of different litter types in arid soils, the activities of lignocellulose-degrading enzymes and respiration were more affected by the changes of litter moisture content than by succession (Fioretto et al. 2007; Papa et al. 2008).

4.6

Conclusions

While there is sufficient information on the most important factors affecting enzymatic activity in forest soils, there are still many questions open for further research. The factors affecting the microscale spatial distribution of enzymes are only partly known and the data regarding the short-time dynamics of enzymatic processes (e.g., after the rewetting of soil) have not been addressed. Also, most of the data so far on the microbial community composition in a soil profile and the succession of biota on decomposing litter are based only on rough estimations of biomass and analyses of culturable microorganisms. However, to link the composition of the enzyme pool in litter with the active portion of the microbial population, it is necessary to simultaneously look for expressed genes and to analyze the total microbial populations using molecular methods. There is also a lack of knowledge about the biochemical properties of soil enzymes, since these enzymes have only scarcely been isolated (Baldrian 2006; Baldrian and Vala´sˇkova´ 2008). Acknowledgments Financial support from the Czech Science Foundation (526/08/0751) and from the Ministry of Education, Youth and Sports of the Czech Republic (OC08050) is gratefully acknowledged.

References Allison SD, Vitousek PM (2004) Extracellular enzyme activities and carbon chemistry as drivers of tropical plant litter decomposition. Biotropica 36:285–296 Allison SD, Vitousek PM (2005) Responses of extracellular enzymes to simple and complex nutrient inputs. Soil Biol Biochem 37:937–944

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Allison SD, Nielsen C, Hughes RF (2006) Elevated enzyme activities in soils under the invasive nitrogen-fixing tree Falcataria moluccana. Soil Biol Biochem 38:1537–1544 Allison SD, Czimczik CI, Treseder KK (2008) Microbial activity and soil respiration under nitrogen addition in Alaskan boreal forest. Global Change Biol 14:1156–1168 Amador JA, Glucksman AM, Lyons JB, Gorres JH (1997) Spatial distribution of soil phosphatase activity within a riparian forest. Soil Sci 162:808–825 Andersson M, Kjoller A, Struwe S (2004) Microbial enzyme activities in leaf litter, humus and mineral soil layers of European forests. Soil Biol Biochem 36:1527–1537 Aneja MK, Sharma S, Fleischmann F, Stich S, Heller W, Bahnweg G, Munch JC, Schloter M (2006) Microbial colonization of beech and spruce litter – influence of decomposition site and plant litter species on the diversity of microbial community. Microb Ecol 52:127–135 Baldrian P (2006) Fungal laccases – occurrence and properties. FEMS Microbiol Rev 30: 215–242 Baldrian P (2008) Enzymes of saprotrophic Basidiomycetes. In: Boddy L, Frankland JC, van West P (eds) Ecology of saprotrophic Basidiomycetes. Academic, Amsterdam, pp 19–41 Baldrian P, Vala´sˇkova´ V (2008) Degradation of cellulose by basidiomycetous fungi. FEMS Microbiol Rev 32:501–521 Baldrian P, Tr€ogl J, Frouz J, Sˇnajdr J, Vala´sˇkova´ V, Merhautova´ V, Cajthaml T, Herinkova´ J (2008) Enzyme activities and microbial biomass in topsoil layer during spontaneous succession in spoil heaps after brown coal mining. Soil Biol Biochem 40:2107–2115 Bastida F, Barbera GG, Garcia C, Hernandez T (2008) Influence of orientation, vegetation and season on soil microbial and biochemical characteristics under semiarid conditions. Appl Soil Ecol 38:62–70 Baum C, Hrynkiewicz K (2006) Clonal and seasonal shifts in communities of saprotrophic microfungi and soil enzyme activities in the mycorrhizosphere of Salix spp. J Plant Nutr Soil Sci 169:481–487 Bengtson P, Falkengren-Grerup U, Bengtsson G (2005) Relieving substrate limitation-soil moisture and temperature determine gross N transformation rates. Oikos 111:81–90 Caldwell BA (2005) Enzyme activities as a component of soil biodiversity: a review. Pedobiologia 49:637–644 Carneiro MAC, Siqueira JO, Moreira FMD, Soares ALL (2008) Soil organic carbon, total nitrogen, microbial biomass and activity in two rehabilitation chronosequences after bauxite mining. Revista Brasileira De Ciencia Do Solo 32:621–632 Carreiro MM, Sinsabaugh RL, Repert DA, Parkhurst DF (2000) Microbial enzyme shifts explain litter decay responses to simulated nitrogen deposition. Ecology 81:2359–2365 Chen HJ (2003) Phosphatase activity and P fractions in soils of an 18-year-old Chinese fir (Cunninghamia lanceolata) plantation. For Ecol Manage 178:301–310 Criquet S, Farnet AM, Tagger S, Le Petit J (2000) Annual variations of phenoloxidase activities in an evergreen oak litter: influence of certain biotic and abiotic factors. Soil Biol Biochem 32:1505–1513 Criquet S, Tagger S, Vogt G, Le Petit J (2002) Endoglucanase and beta-glycosidase activities in an evergreen oak litter: annual variation and regulating factors. Soil Biol Biochem 34: 1111–1120 Criquet S, Ferre E, Farnet AM, Le Petit J (2004) Annual dynamics of phosphatase activities in an evergreen oak litter: influence of biotic and abiotic factors. Soil Biol Biochem 36:1111–1118 Dilly O, Munch JC (1996) Microbial biomass content, basal respiration and enzyme activities during the course of decomposition of leaf litter in a black alder (Alnus glutinosa (L) Gaertn) forest. Soil Biol Biochem 28:1073–1081 Dong S, Brooks D, Jones MD, Grayston SJ (2007) A method for linking in situ activities of hydrolytic enzymes to associated organisms in forest soils. Soil Biol Biochem 39:2414–2419 Fioretto A, Papa S, Curcio E, Sorrentino G, Fuggi A (2000) Enzyme dynamics on decomposing leaf litter of Cistus incanus and Myrtus communis in a Mediterranean ecosystem. Soil Biol Biochem 32:1847–1855

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Fioretto A, Papa S, Pellegrino A, Fuggi A (2007) Decomposition dynamics of Myrtus communis and Quercus ilex leaf litter: mass loss, microbial activity and quality change. Appl Soil Ecol 36:32–40 G€om€oryova´ E (2004) Small-scale variation of microbial activities in a forest soil under a beech (Fagus sylvatica L.) stand. Pol J Ecol 52:311–321 G€om€oryova´ E, Gregor J, Pichler V, G€ om€ ory D (2006) Spatial patterns of soil microbial characteristics and soil moisture in a natural beech forest. Biologia 61:S329–S333 Gramss G (1997) Activity of oxidative enzymes in fungal mycelia from grassland and forest soils. J Basic Microbiol 37:407–423 Kahkonen MA, Wittmann C, Kurola J, Ilvesniemi H, Salkinoja-Salonen MS (2001) Microbial activity of boreal forest soil in a cold climate. Boreal Environ Res 6:19–28 Kjoller A, Struwe S (1990) Decomposition of beech litter: a comparison of fungi isolated on nutrient rich and nutrient poor media. Trans Mycol Soc Jpn 31:5–16 Kjoller AH, Struwe S (2002) Fungal communities, succession, enzymes, and decomposition. In: Burns RG, Dick RP (eds) Enzymes in the environment. Marcel Dekker, New York, Basel, pp 267–284 Kramer S, Green DM (2000) Acid and alkaline phosphatase dynamics and their relationship to soil microclimate in semiarid woodland. Soil Biol Biochem 32:179–188 Landgraf D, Wedig S, Klose S (2005) Medium- and short-term available organic matter, microbial biomass, and enzyme activities in soils under Pinus sylvestris L. and Robinia pseudoacacia L. in a sandy soil in NE Saxony, Germany. J Plant Nutr Soil Sci 168:193–201 Linkins AE, Sinsabaugh RL, McClaugherty CA, Melills JM (1990a) Cellulase activity on decomposing leaf litter in microcosms. Plant Soil 123:17–25 Linkins AE, Sinsabaugh RL, McClaugherty CA, Melillo JM (1990b) Comparison of cellulase activity on decomposing leaves in a hardwood forest and woodland stream. Soil Biol Biochem 22:423–425 Luis P, Kellner H, Zimdars B, Langer U, Martin F, Buscot F (2005) Patchiness and spatial distribution of laccase genes of ectomycorrhizal, saprotrophic, and unknown basidiomycetes in the upper horizons of a mixed forest cambisol. Microb Ecol 50:570–579 Moller J, Miller M, Kjoller A (1999) Fungal–bacterial interaction on beech leaves: influence on decomposition and dissolved organic carbon quality. Soil Biol Biochem 31:367–374 Morris SJ (1999) Spatial distribution of fungal and bacterial biomass in southern Ohio hardwood forest soils: fine scale variability and microscale patterns. Soil Biol Biochem 31:1375–1386 Mosca E, Montecchio L, Scattolin L, Garbaye J (2007) Enzymatic activities of three ectomycorrhizal types of Quercus robur L. in relation to tree decline and thinning. Soil Biol Biochem 39:2897–2904 M€ ott€onen M, J€arvinen E, Hokkanen TJ, Kuuluvainen T, Ohtonen R (1999) Spatial distribution of soil ergosterol in the organic layer of a mature Scots pine (Pinus sylvestris L.) forest. Soil Biol Biochem 31:503–516 Niemi RM, Vepsalainen M, Erkomaa K, Ilvesniemi H (2007) Microbial activity during summer in humus layers under Pinus sylvestris and Alnus incana. For Ecol Manage 242:314–323 Olander LP, Vitousek PM (2000) Regulation of soil phosphatase and chitinase activity by N and P availability. Biogeochemistry 49:175–190 Orchard VA, Cook FJ (1983) Relationship between soil respiration and soil moisture. Soil Biol Biochem 15:447–453 Osono T (2007) Ecology of ligninolytic fungi associated with leaf litter decomposition. Ecol Res 22:955–974 Osono T, Takeda H (2005) Decomposition of organic chemical components in relation to nitrogen dynamics in leaf litter of 14 tree species in a cool temperate forest. Ecol Res 20:41–49 Papa S, Pellegrino A, Fioretto A (2008) Microbial activity and quality changes during decomposition of Quercus ilex leaf litter in three Mediterranean woods. Appl Soil Ecol 40:401–410 Prietzel J (2001) Arylsulfatase activities in soils of the Black Forest/Germany – seasonal variation and effect of (NH4)2SO4 fertilization. Soil Biol Biochem 33:1317–1328

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Renella G, Landi L, Valori F, Nannipieri P (2007) Microbial and hydrolase activity after release of low molecular weight organic compounds by a model root surface in a clayey and a sandy soil. Appl Soil Ecol 36:124–129 Ross DJ (1987) Soil microbial biomass estimated by the fumigation-incubation procedure: seasonal fluctuations and influence of soil moisture content. Soil Biol Biochem 19:397–404 Saetre P (1999) Spatial patterns of ground vegetation, soil microbial biomass and activity in a mixed spruce-birch stand. Ecography 22:183–192 Saetre P, Baath E (2000) Spatial variation and patterns of soil microbial community structure in a mixed spruce-birch stand. Soil Biol Biochem 32:909–917 Saiya-Cork KR, Sinsabaugh RL, Zak DR (2002) The effects of long term nitrogen deposition on extracellular enzyme activity in an Acer saccharum forest soil. Soil Biol Biochem 34:1309–1315 Sardans J, Penuelas J (2005) Drought decreases soil enzyme activity in a Mediterranean Quercus ilex L. forest. Soil Biol Biochem 37:455–461 Scheu S, Wirth S, Eberhardt U (1993) Decomposition of C-14-labeled cellulose substrates in litter and soil from a beechwood on limestone. Microb Ecol 25:287–304 Schlatte G, Kampichler C, Kandeler E (1998) Do soil microarthropods influence microbial biomass and activity in spruce forest litter. Pedobiologia 42:205–214 Selmants PC, Hart SC, Boyle SI, Stark JM (2005) Red alder (Alnus rubra) alters community-level soil microbial function in conifer forests of the Pacific Northwest, USA. Soil Biol Biochem 37:1860–1868 Sinsabaugh RL, Carreiro MM, Alvarez S (2002a) Enzyme and microbial dynamics of litter decomposition. In: Burns RG, Dick RP (eds) Enzymes in the environment. Marcel Dekker, New York, Basel, pp 249–265 Sinsabaugh RL, Carreiro MM, Repert DA (2002b) Allocation of extracellular enzymatic activity in relation to litter composition, N deposition, and mass loss. Biogeochemistry 60:1–24 Sˇnajdr J, Vala´sˇkova´ V, Merhautova´ V, Cajthaml T, Baldrian P (2008a) Activity and spatial distribution of lignocellulose-degrading enzymes during forest soil colonization by saprotrophic basidiomycetes. Enzyme Microb Technol 43:186–192 Sˇnajdr J, Vala´sˇkova´ V, Merhautova´ V, Herinkova´ J, Cajthaml T, Baldrian P (2008b) Spatial variability of enzyme activities and microbial biomass in the upper layers of Quercus petraea forest soil. Soil Biol Biochem 40:2068–2075 Stoyan H, De-Polli H, Bohm S, Robertson GP, Paul EA (2000) Spatial heterogeneity of soil respiration and related properties at the plant scale. Plant Soil 222:203–214 Timonen S, Sen R (1998) Heterogeneity of fungal and plant enzyme expression in intact Scots pine Suillus bovinus and Paxillus involutus mycorrhizospheres developed in natural forest humus. New Phytol 138:355–366 Trasar-Cepeda C, Leiros MC, Gil-Sotres F (2000) Biochemical properties of acid soils under climax vegetation (Atlantic oakwood) in an area of the European temperate-humid zone (Galicia, NW Spain): specific parameters. Soil Biol Biochem 32:747–755 van der Wal A, van Veen JA, Smant W, Boschker HTS, Bloem J, Kardol P, van der Putten WH, de Boer W (2006) Fungal biomass development in a chronosequence of land abandonment. Soil Biol Biochem 38:51–60 Venkatesan S, Senthurpandian VK (2006) Comparison of enzyme activity with depth under tea plantations and forested sites in south India. Geoderma 137:212–216 Waldrop MP, Zak DR (2006) Response of oxidative enzyme activities to nitrogen deposition affects soil concentrations of dissolved organic carbon. Ecosystems 9:921–933 Waldrop MP, McColl JG, Powers RF (2003) Effects of forest post harvest management practices on enzyme activities in decomposing litter. Soil Sci Soc Am J 67:1250–1256 Wittmann C, Kahkonen MA, Ilvesniemi H, Kurola J, Salkinoja-Salonen MS (2004) Areal activities and stratification of hydrolytic enzymes involved in the biochemical cycles of carbon, nitrogen, sulphur and phosphorus in podsolized boreal forest soils. Soil Biol Biochem 36:425–433

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Zhang YM, Zhou GY, Wu N, Bao WK (2004) Soil enzyme activity changes in different-aged spruce forests of the eastern Qinghai-Tibetan plateau. Pedosphere 14:305–312 Zhang YM, Wu N, Zhou GY, Bao WK (2005) Changes in enzyme activities of spruce (Picea balfouriana) forest soil as related to burning in the eastern Qinghai-Tibetan Plateau. Appl Soil Ecol 30:215–225

.

Chapter 5

Extracellular Enzymes in Sensing Environmental Nutrients and Ecosystem Changes: Ligand Mediation in Organic Phosphorus Cycling Thanh H. Dao

5.1

Introduction

Phosphorus (P) is a nonmetallic element that is essential to living organisms. Plant available P is growth limiting and organic P contributes to plant mineral nutrition only after dephosphorylation and release of inorganic phosphate, which is of particular importance in phosphate-deficient natural or cultivated ecosystems. Most of the organic P mineralizing activity occurs in the rhizosphere of plants where extracellular phosphohydrolases are released by plant roots (Tanaka et al. 1974; Helal 1990; Duff et al. 1994; Hunter and McManus 1999; Lopez-Gutierrez et al. 2004; Ma et al. 2009) and microorganisms (Greiner et al. 2000; Quan et al. 2003; Vohra and Satyanarayana 2003; Dao and Hoang 2008), allowing these organisms access to outlying P substrates that otherwise are not accessible (Li et al. 1991; Joner and Johansen 2000; Ezawa et al. 2005). However, multiple competing reactions are operating to regulate the solution-phase concentration and availability of these P-containing substrates and the released phosphate for plant uptake or dispersion in the environment (Fig. 5.1). Thus, to gain an accurate description of the transformations and cycling of organic P in soils and the environment, it is important to understand the interactions that occur between plant and their roots, microorganisms and fauna that populate terrestrial and aquatic environments, and the processes that are involved in the cycling of P-compounds in those biospheres. In this chapter, we will give particular attention to the ligand exchange process involved in the release of stabilized organic P, a forerunner process crucial to the action of phosphohydrolases. The question of why these enzymes are extracellular has always been of intense interest. It is often wondered why plants and microorganisms externalize low quantities of such entities into their surrounding where the risks of sorption, deactivation,

T.H. Dao US Department of Agriculture, Agriculture Research Service, Beltsville, MD, USA e-mail: [email protected]

G. Shukla and A. Varma (eds.), Soil Enzymology, Soil Biology 22, DOI 10.1007/978-3-642-14225-3_5, # Springer-Verlag Berlin Heidelberg 2011

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Fig. 5.1 Processes and reactions affecting phosphorus pools in soil

or degradation are relatively high in soil and aquatic environments (Burns and Dick 2002). Burns (1982) postulated that the localization of metabolic enzymes was partially determined by the size and the solubility of the target substrates, and the enzymes’ activity regulated by whether they freely move in the soil or are in association with a mineral or organic surface of soil colloids. Although they may be short-lived, the extracellular enzymes are responsible for the hydrolysis of organic substrates that cannot pass through cell walls or membranes without extensive expenditures of cell’s resources or that the substrate molecules are simply too large to do so. Phosphohydrolases are synthesized to search out P-containing compounds and generate signals to which the producer’s cells react by increasing expression and production of additional enzymes. Although enzymes’ activity may be attenuated (Dick and Tabatabai 1987; Quiquampoix 1987; Quiquampoix et al. 2002; George et al. 2005), the apparent risk to externalizing enzymes is minimized by these surface interactions and associations (Burns 1982). A highly effective strategy of the cell, from a root of a plant, or an entire microbial community is to have a group of catabolic enzymes associated at or near the cell’s surface, performing a sequence of processes in close proximity; an ideal biochemical microenvironment is thereby created, where complementary functions and metabolite concentrations are much higher than those of the surrounding to maximize the efficacy of individual metabolic processes. As plant and microorganisms have evolved multiple strategies

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to obtain the needed P, it is not surprising to observe the interweaving of physical and biochemical processes in the speciation and availability of organic P forms. Therefore, in our efforts to gain a better understanding of the processes of P sensing and cycling, a combination of ligand exchange and mineralization of organic P in an analytical tool for determining organic nutrient turnover or for detecting ecosystem changes appears to be a natural blend (Dao 2003, 2004, 2007; Bunemann 2008). From an environmental standpoint, the contribution of biologically mediated processes to the organic P release and movement must be fully understood to develop sustainable management practices for our agricultural production systems and mitigate eutrophic responses in sensitive aquatic ecosystems. In spite of major research and education efforts and public media scrutiny, agricultural P continues to be a significant pollutant of surface waters and estuaries around the globe. As soil fertility and nutrient accumulation rise, P-enriched soils remain a major contributor to P loading to runoff and surface and ground water as the result of land application of fertilizers and organic nutrients (Council for Agricultural Science and Technology CAST 1996, 2002; Zhang et al. 2006; Green et al. 2007; Dao et al. 2008). Algal blooms have often been related to increased P inputs from terrestrial ecosystems, often exceeding a critical soluble P concentration of 30 mg L
1 (Smith 1982; Jarvie et al. 2002; Mainstone and Parr 2002; Haukka et al. 2006; Hilton et al. 2006). Therefore, we will take another fundamental look at the biogeochemistry of P in the soil to understand the solubilization of organic P substrates, and the role of ligands in the enzyme-mediated mineralization and turnover of organic P in the supply of phosphate to meet plant and microbial nutritional needs.

5.2

Biogeochemistry of Phosphorus

Phosphorus is a decay product of Cl, S, and K (Lal et al. 1988). Radioactive decay series of 34Ca, 33K, 32K, 31F, 33Ne, and 34Ne isotopes have 31P as a daughter product via beta and proton emissions. Phosphorus is always found combined with other elements, particularly oxygen. In a tetrahedral arrangement, four oxygen atoms bond to a P atom to form the phosphate (PO43
) anion. Each oxygen atom carries a negative charge and can bond with hydrogen to yield, HPO42
, H2 PO4
, and H3PO4, or phosphoric (V) acid, a major industrial product for the fertilizer and food industries. Phosphorus also can have an oxidation state of þ3, phosphorus (III) which yields a large group of important industrial compounds and pesticides, including PH3, PCl3, P2S5, and phosphonates. The chemistry and metabolism of reduced organic P compounds have been extensively reviewed elsewhere (Devai et al. 1988; Ternan et al. 1998; Roels and Verstraete 2001; Quinn et al. 2007; White and Metcalfe 2007). Phosphorus circulates through the environment in three natural cycles. In the inorganic cycle, phosphate-P in the earth crust is released to the soil, passes to lakes, streams and rivers to the seas and ocean floors. Under geologic pressure and temperature, P-containing sediments undergo transformations, resulting in

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metamorphic rocks that may be uplifted to form new landmasses to begin again the cycle, from land to sea floors. Phosphate rock is widely distributed across the globe in marine phosphorite deposits. Apatite, i.e., calcium phosphates, is the primary P compound in phosphate rock. Phosphorus in apatite minerals is only slightly soluble and of limited availability to plants and must be chemically processed to produce many agriculturally important forms of P fertilizers used in crop production, such as superphosphates, triple superphosphate, or ammonium phosphates when combined with ammonia. Superimposed upon the inorganic P cycle, the other two cycles are about the moving of P through living organisms that are either aquatic or terrestrial dwellers. In these organisms, P is a required nutrient, playing a central role in numerous energetic metabolic processes. For example, P is a component of phospholipids, nucleic acids, the principal nucleotide cofactors required for energy transfer and catalysis in the cell, or in inositol phosphates involved in signal transduction and cellular regulation. Therefore, most P-containing compounds found in living systems are in the form of inorganic phosphate and organic phosphate mono- and diesters. The latter compounds are characterized by a carbon–oxygen–phosphorus bond, where P has a valence of þ5. The biochemistry of P-containing compounds is considered relatively simple, consisting almost entirely of phosphate ester and phospho-anhydride formation and hydrolysis.

5.2.1

Inorganic Phosphate

Phosphate may undergo complex reactions with soil mineral and organic constituents of natural waters and soils. In dilute solutions, the following species are in thermodynamic equilibrium (Table 5.1). Inorganic phosphate (i.e., orthophosphate) is a reactive chemical species, which is rapidly incorporated in various soil P pools, making most of the applied P unavailable for plant uptake. As a case study, Lindsay and Stephenson (1959) made a detailed study of the reactions affecting a granule of monocalcium phosphate monohydrate [Ca(H2PO4)2H2O], a common superphosphate fertilizer, when added to soil. Upon contact with the soil, the granule undergoes a series of complex reactions that includes dissolution and precipitation to transform it into more stable P minerals that render P even less available to plants. In sequence, a Table 5.1 Dissociation constants for various phosphate species

H3PO40 Ð H+ + H2PO4
H2PO4
Ð H+ + HPO42
HPO42
Ð H+ + PO43
H2PO4
Ð 2H+ + PO43
2H2PO4
Ð (H2PO4)22
Lindsay (1979)

log K
2.15
7.20
12.35
19.55
0.35

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supersaturated solution of Ca2+, H2PO4
, CaHPO42H2O, and CaHPO4 is formed at the water-fertilizer granule boundary. Super saturation induces the precipitation in the intermediary metastable solution of varying proportion of the two minerals, CaHPO42H2O and CaHPO4 (Lindsay and Stephenson 1959). A sharp pH decrease also occurs as H3PO40 is formed. The highly acidic solution reacts with soil mineral surfaces, releasing Al3þ, Fe3þ, Fe2þ, Mn2þ, and other cations forming aquo-metal ions such as Al(H2O)63þ or Fe(H2O)63þ; some of the acidity is neutralized in the process and initial reaction products may include aluminum and iron phosphates. Upon the addition of Ca(H2PO4)2H2O to three soils, these researchers have identified some 32 aluminum ferric phosphate minerals of variable composition, (Al, Fe, X)-PO4nH2O where X denotes the presence of cations other than Al and Fe (Lindsay et al. 1962). In addition to reactions in the solution phase, Henmi and Huang (1985) demonstrated strong sorption of phosphate by edge surface AlO-functional groups in soils developed from volcanic ash and pumice. Components in Andisols, which are responsible for adsorption of P, are amorphous or poorly ordered hydrous aluminosilicates in the clay fraction, namely, allophanes and imogolite. Grinding or heating the clays at temperatures between 200 and 400 C increased P sorption. The increased capacity was attributed to the formation of the new P adsorption sites (AlOH–H2O), as the result of breaking down Si–O–Al linkages located only on the edge surface of allophane and imogolite tubular structures in the clay fraction. In acid soils, phosphate strong sorption has been attributed to hydrous oxides of Fe and Al, and to (1:1) layer lattice clays (Juo and Fox 1977; Golberg and Sposito 1984b; Sanyal et al. 1993; Arai and Sparks 2002). In calcareous soils, the abundance of Ca and Mg results in the sorption and precipitation of pyrophosphates as insoluble compounds such as Ca(NH4)2P2O7H2O and Mg(NH4)2P2O74H2O, and in the process, reduces the amount of phosphate remaining in the soil solution for uptake by microorganisms or plant roots (Lindsay et al. 1962). Phosphate adsorption on soils often has been described mathematically with the following generalized equation: qðCÞ ¼

n X i¼1



bi Kibi Cbi 1 þ Kibi Cbi



where q(C) is the quantity of phosphate adsorbed per unit mass of soil solids, C is the solution- phase concentration of phosphate, and n, bi, Ki, and bi, are empirical parameters (Golberg and Sposito 1984). The equation takes the familiar form of the Langmuir adsorption model, x/m ¼ Smax (KC/(1 þ KC), when n ¼ 1 and bi ¼ 1; and the equation reduces to the Freundlich equation, S ¼ KCn, when n ¼ 1, 0 < bi <1, and KC << 1. Goldberg and Sposito (1984a) reported that the constant capacitance model developed by Schindler et al. (1976) and Stumm et al. (1976) provides more appropriately a quantitative description of phosphate adsorption by a variety of aluminum and iron hydrous oxides. The model describes phosphate adsorption

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based on a ligand exchange mechanism, and relates the charge density of a surface or plane of charge (so) to the surface potential (Co), capacitance density C (farad m
2), surface area, S (m2 g
1), the Faraday constant, F (coulomb mol
1), and the suspension density (g L
1). Thus, diverse reactions and transformations affect inorganic phosphates in the soil, illustrating the complexity of the chemical environment which plant roots and microorganisms face in their quest for P from their growth medium.

5.2.2

Organic Phosphorus

In terrestrial and aquatic ecosystems, microorganisms have evolved the ability to store a number of necessary metabolites in the form of polymers that are osmotically inert. Inorganic phosphate is one such metabolite that is stored within the cell as a high-molecular-weight inorganic polyphosphate; it is a linear polymer containing from a few to several hundred residues of phosphate that are linked by energyrich phosphoanhydride bonds (Kulaev and Kulakovskaya 2000). In microbial cells, polyphosphate can confer the organism the increased resistance to stressful changes in its environment, regulation of enzyme activities, cation transport, as well as a source of inorganic phosphate to meet energy needs. Another important group of stored phosphorylated organic compounds include inositol phosphate monoesters and phosphate diesters (i.e., phospholipids and nucleic acids). Plant reproductive organs and, in particular, seeds of wheat (Triticum aestivum L.), rice (Oriza sativa L.), corn (Zea mays L.), soybean [Glycine max (L.) Merr.], and mung bean [Vigna radiata (L.) R. Wilczek var. radiata] (Asada et al. 1969; Tanaka et al. 1974; Yamagata et al. 1980; Scott and Loewus 1986; Lott et al. 2000; Shi et al. 2005) store P as calcium, magnesium, and potassium salts of myo-inositol hexakis dihydrogenphosphate (hereafter, will be referred to as myo-inositol hexakisphosphate). These compounds supply inositol and a range of phosphorylated substrates, inositol as C source, and cations to the growing embryo upon seed germination. The chemistry of inositol phosphate monoesters is found extensively in the recent literature due to their expansive roles in biological systems (e.g., cellular signal transduction; calcium regulation) and in plants and soil organisms (e.g., condensed P storage; antioxidant) (Cosgrove 1980; Stewart and Tiessen 1987; Voglmaier et al. 1996; Kulaev and Kulakovskaya 2000; Lemtiri-Chlieh et al. 2003; Turner et al. 2005, 2007). The turnover of microbial tissues and plant residues at the soil surface contributes organic P compounds to the soil environment (Anderson 1980; Stewart and Tiessen 1987). Cultural practices such as the incorporation of green cover crops into the soil, and land application of animal manure also add plant-derived P to the soil P pools in an effort to build up soil fertility and thereby productivity (Biederbeck et al. 1993; Cavigelli and Thien 2003). myo-Inositol hexakisphosphate and lower phosphomonoesters of inositol are the most abundant organic P compounds in soils, and may make up to 50% of total soil organic P (Wild and Oake 1966; Koopmans et al. 2003; Dao 2004). Of all the forms of organic P, the most prevalent

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forms found in soil are several steroisomers of inositol hexakisphosphate as they are recalcitrant to microbial degradation. These phosphomonoesters have been identified using 31P nuclear magnetic resonance spectroscopy (Cade-Menun et al. 2002; Koopmans et al. 2003; Turner et al. 2003; Jayasundera et al. 2005), high-performance liquid chromatography and mass spectrometry (Sandberg and Ahderinne 1986; Lehrfeld 1989, 1994; Cooper et al. 2007). Information on the nature of organic P in manure is also increasingly available for different kinds of livestock and poultry and manure management practices (Peperzak et al. 1959; Gerritse and Eksteen 1978; He and Honeycutt 2001; Dao et al. 2006). As manure is composed of partly digested feed, direct spectroscopic measurements showed that the organic P fraction generally consists of phosphomonoesters of inositol, phospholipids, and nucleic acids derived from plant and microbial materials. In soils amended with animal and green manure from cover crops and plant residues, phytic acid (myo-inositol hexakisphosphate) and other organic P are added to the near-surface soil P pool, in addition to inorganic phosphate (Dao 2004; Zhang et al. 2006). Although there have been many studies on the behavior of inorganic phosphate in soil as discussed in a previous section, desorption, speciation, and mineralization of organic P compounds is yet poorly understood. Mechanistic studies of the interactions between specific organic P compounds, metal hydrous oxides, and soil mineral and organic colloids as affected by solution properties such as pH, electrical conductivity, or other cation concentration are sparse.

5.3

Reactions Involving Organic Phosphorus in Soils

5.3.1

Chemical Processes

5.3.1.1

Adsorption–Precipitation

Although mathematically and mechanistically are very similar, differentiation between adsorption and precipitation is at the soil–liquid interface (Sposito 1986; Lutzenkirchen and Behra 1996). Many of the solute–surface interactions, namely, absorption, adsorption or enrichment at the interface of the solution and solid soil constituents, diffusion into the soil solids, and precipitation or the formation of a P solid phase at the interface have often been referred to as sorption when most if not all these mechanisms are operating to control P speciation and environmental behavior. The mechanisms of of phosphate sorption on soils have been extensively studied; however, those of organic phosphate monoesters or organic P in general, have not received the same degree of scientific scrutiny. As they are biochemically reactive, interacting with soil components stabilize and shield phosphate monoesters from biodegradation, leading to their preferential accumulation in soils

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(Anderson et al. 1974; Stewart and Tiessen 1987; Celi and Barberis 2007). The adsorption of phosphate monoesters on hydrous oxides has been modeled on inorganic phosphate adsorption on iron hydrous oxides (Ognalaga et al. 1994; Celi et al. 2001). Ognalaga et al. (1994) advanced a model of a binuclear surface complex for the adsorption of myo-inositol hexakisphosphate on dried samples of goethite that was equilibrated with the inositol hexakisphosphate (Fig. 5.2). In addition, whether the ring structure was myo-inositol or glucose, these researchers observed that the organic portion of the molecule did not appear to have any effect on the adsorption process, upon comparing the sorption of glucose-1-phosphate and myo-inositol hexakisphosphate onto goethite. This hypothesis was derived from the model of the adsorbed state of inorganic phosphate on ferrihydrite and other iron oxides (Hingston et al. 1974; Parfitt 1979; Parfitt et al. 1975) and soil constituents (Parfitt 1977). By ligand exchange, two surface hydroxyl ions (or water molecules) were replaced by a phosphate ion. Two of the oxygen atoms of the phosphate ion were coordinated to two different Fe3þ ions, resulting in a binuclear surface complex of the type Fe
O
P(O2)
O
Fe (Hingston et al. 1974; Parfitt et al. 1975; Beek and Riemsdijk 1982). Molecular cross-sectional areas and surface coverage calculations and X-ray photoelectron spectroscopy further supported the binuclear model of phosphate adsorption (Martin and Smart 1987; Shin et al. 2004). In such an arrangement, the quantity

Fig. 5.2 Proposed adsorbed state of myo-inositol hexakisphosphate on goethite. The boat conformation of the molecule was attributed to Cosgrove DJ 1966. The H
and OH
groups of the phosphate moieties are not shown (Adapted from Ognalaga et al. (1994))

5 Extracellular Enzymes in Sensing Environmental Nutrients

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of adsorbed phosphate per unit surface area averaged 2.5 mmol m
2 for a number of samples of natural and synthetic goethite (Golberg and Sposito 1984; Torrent et al. 1990). Variability in sorption capacities of iron and aluminum hydrous oxides evolve from differences in availability of singly coordinated
OH groups on the mineral surface (Parfitt 1989) (Fig. 5.3). It is noted that, in the binuclear surface complex model for an organic phosphate monoester, myo-inositol hexakisphosphate, the adsorbed molecule is forced into the boat conformation, deviating from its thermodynamically stable chair conformation. Furthermore, formation of monodentate complexes may well be an important mechanism of adsorption of phosphate monoesters in soil-water suspensions or colloids and sediments in aquatic ecosystems, as was the case for phosphate in the presence of water (Golberg and Sposito 1985). Shin et al. (2004) showed that phosphate adsorption occurred at least through a mixture of monodentate and binuclear bridges (Fig. 5.4), using a combination of infrared spectroscopy and X-ray photoelectron and diffraction spectroscopic techniques. In the author’s opinion, one can also visualize binuclear and polynuclear complexes for the sorption of inositol phosphate monoesters, forming monodentate bridges with FeOþ and -AlOþ of more than one surfaces of suspended colloids in soil suspensions and sediments.

5.3.1.2

Autohydrolysis and Metal-Mediated Hydrolysis

Other deactivation reactions involving organic P include autohydrolysis in presence of metal ions. myo-Inositol hexakisphosphate may not be chemically hydrolyzed in vivo in the pH and temperature range of biological systems. However, its autohydrolysis may be triggered by the coordination of the metal ion with a phosphate group of myo-inositol hexakisphosphate, promoting the hydrolysis of an adjacent phosphate group. Bullock et al. (1993) observed its spontaneous chemical hydrolysis at 95  C across a wide range of ambient pH. The reaction is significant in efforts to negate the binding and sequestration of essential trace minerals with the consumption of high-phytate food; boiling such food would destroy its binding and sequestering, for example dietary Fe to interfere with Fe absorption in humans on a high fiber diet (Sandberg et al. 1999). Specific instances of the autohydrolysis of myo-inositol hexakisphosphate have also been observed at high temperature of 120 C, resulting in the formation of most of the phosphate esters of myo-inositol, with 39 chromatographically distinct myo-inositol mono-, bis-, tris-, tetrakis, pentakis-, and hexakisphosphates in the presence of HCl (Hull et al. 1999). Metal-catalyzed dephosphorylation of nucleoside-50 trisphosphates was observed in aqueous solutions by nucleophilic substitution with water in the presence of Cu2+, Zn2+, Ni2+, Mn2+, and Mg2+ (Sigel et al. 1984). Details on SN1 and SN2 nucleophilic substitution mechanisms are found in a review of the topic by Baldwin et al. (2005).

84

T.H. Dao 100 Hematite

1-day 3-day 10-day 30-day 10 1

10

100

10

100

200

Sorbed P concentration (mmol kg–1)

Geothite

100 90 80 70 60 50 40 1 1000 Allophane

100

10 0.1

1

10

1

10

100

1000 Ferrihydrite

100

10 0.1

Solution phase P concentration (umol Fig. 5.3 Sorption of phosphate on aluminum and iron oxides (Parfitt 1989)

100

L–1)

5 Extracellular Enzymes in Sensing Environmental Nutrients

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Fig. 5.4 Possible phosphate surface complexes (Golberg and Sposito 1985)

5.3.2

Biological Processes Involved in Organic P Solubilization and Degradation

As plants can only acquire P as inorganic phosphate, recycling of P has to occur through the turnover of biological tissues and the mineralization of organic P in its environment (Tadano and Sakai. 1991; Frossard et al. 2000; Hinsinger 2001). The process is mediated by phosphohydrolases induced and released by microbial systems, plant roots, or arbuscular mycorrhizal associations to mineralize organic P forms and release phosphate for plant assimilation (Harrison 2005). For example, wheat (Triticum aestivum L.) roots were observed to secrete 5–10 times greater concentrations of phosphohydrolases in the presence of organic P, when compared to a sterilized medium containing only inorganic P (Tarafdar and Claassen 2003). Roots of a wide range of plant species have shown similar ability to meet their P nutritional needs, e.g., white lupin (Lupinus albus L.) (Wasaki et al. 2003) and clover (Hunter and McManus 1999). It thus appears that mediated processes a plant uses to obtain P from its environment have an efficiency that by far exceeds that of abiotic processes to lead to the eventual release of phosphate for bringing the nutrient inside the cell via an active or passive process across its membrane (Ragothama 1999; Horst and Kahm 2004). Phosphate transporters in plant roots and cultured microbial cells are numerous and have been extensively studied (Ragothama 1999; Kulaev and Kulakovskaya 2000). Transporters are proteins that are an integral component of the cell membrane, capable of internalizing inorganic phosphate and other simple molecules; the transport is based on a chemiosmotic ion gradient and an example of a phosphate transporter in Arabidopsis thaliana has been described in Raghothama’s review article (1999). Following is another basic rendition of the phosphate transporter interacting with organic anions in the rhizosphere by Horst and Kahm (2004) (Fig. 5.5). 5.3.2.1

Mineralization of Environmental Biological Tissues Containing Organic P

In native ecosystems, growing plants depend upon the mineralization of P-containing organic tissues and plant residues to meet their biotic needs, particularly in phosphate-deficient ecosystems. Suites of catabolic enzymes (proteases, glucosidases,

86

T.H. Dao H2-PO4–

R-COO– ADP + P1

ATP

H+ - ATPase H+

H+ H+

ch

d

n ga

om

C

n tio lu so

is

Microbial biomass

ion

pt

or

s De

n io at ex pl

n

io

et

cr

Ex

H2-PO4–

Fe D

n

tio

da

ra

g De

ex

Li

R-COO–

transport

e

g an

m

Co

R-COO– - Fe - H2 -PO4–

n

io

at

ex

pl

n

tio

rp

so

Fe/AI oxides

Fig. 5.5 Ligand-mediated phosphate release and transport across plant roots’ membrane (Adapted from Horst and Kahm (2004))

oxidases, and phosphohydrolases) are induced and released to decompose organic matter and obtain the organic P contained within. Although much of the literature on the decomposition of plant residues and forest litter has been focused on the mineralization of C and N (Douglas et al. 1980; Christensen 1986; Bremer and van Kessel 1992; Dao 1998), significant interactions were observed between P and the mineralization of plant residue C and N (Kouno et al. 2002; Soon and Arshad 2002; Dao and Cavigelli 2003; Lupwayi et al. 2007). Mineralization of P was found to occur with substrates having C:P ratios near 30–50 though most plant residues have C:P rations exceeding 140:1.

5.3.2.2

Mycorrhizal Associations

To increase the efficiency of the scavenging for P-containing substrates, symbiotic association between a host plant and certain soil fungi, known as arbuscular mycorrhiza (AM) have been part of plant strategies to fulfill their P needs in natural and cultivated ecosystems. The arbuscular mycorrhizae (AM) hyphae can explore a large volume of soil scavenging for P substrates, inducing physiological changes that favor the establishment of P-mineralizing microorganisms in the hyphal microenvironment (Dighton 1983, 1991; Tarafdar and Marschner 1994; Singh and Kapoor 1998; Lopez-Gutierrez et al. 2004). Numerous studies have shown that mycorrhizae dissolve insoluble P minerals (Reid et al. 1984; Leyval and Reid 1991) through the excretion of H+ and organic acids (Li et al. 1991; Lapeyrie et al. 1991; Neumann and Martinoia 2002), and hydrolysis of organic P by excreted phosphohydrolases. The contributions of AM to the cycling of P have been periodically

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reviewed (Bonfante-Fasolo 1984; Smith and Gianinazzi-Pearson 1988; Harrison 2005) and will not be further discussed in this chapter.

5.3.2.3

Extracellular Phosphohydrolases

Common to the above two mechanisms for obtaining P from their environment, microorganisms and plants are known to produce and release phosphohydrolases to the external environment, allowing the plant roots access to the outlying P-containing substrates otherwise inaccessible to the host plant (Li et al. 1991; Tadano and Sakai 1991; Jayachandran et al. 1992; Joner and Johansen 2000). By far, the biological dephosphorylation of inositol phosphates is the prevalent pathway of their breakdown, and is carried out by extracellular phosphohydrolases. Histidine acid- and alkaline phosphohydrolases are produced by a wide variety of organisms, including bacteria (i.e., Bacillus subtilis), yeasts (i.e., Saccharomyces), and fungi (i.e., Aspergillus) (Vohra and Satyanarayana 2003). Extensive reviews of organisms capable of utilizing inositol phosphates have been made (Greiner 2007). These enzymes are industrially produced as animal feed supplements (Vats et al. 2005). Many current developments include efforts to improve phosphohydrolases’ activity at animal gastro-intestinal pH, their thermal stability and substrate specificity (Weaver et al. 2007). From an environmental standpoint, the contribution of biologically-mediated processes to organic P release and movement must be fully understood to develop management practices to mitigate P loading to surface waters. Runoff from P-enriched cropping systems showed elevated P concentration (Green et al. 2007; Dao et al. 2008). While run off from P-enriched soils contains more P than the functionally defined molybdate-reactive P forms, information on the biological significance of the remaining unreactive fraction is limited. A study was conducted to characterize distributions of inorganic and enzyme-labile P forms in simulated runoff from soils under an orchardgrass-red clover and a no-till forage-type soybean-winter wheat systems showed significant relationships between waterextractable P (WEP) fluxes and soil bioactive P pools. Concentration and mass distributions of P forms in runoff over time were log-normally distributed. The strength of correlations between cumulative WEP mass loads and soil P pools was in the decreasing order of ligand-exchangeable organic P > ligand-exchangeable inorganic P > Mehlich-3 extractable P, suggesting that runoff P forms were directly associated with soil available P fractions that were partly derived from enzyme-mediated processes.

Enzymatic Dephosphorylation of Uncomplexed Organic Phosphate Monoesters Stored organic P in seeds in the forms of calcium and magnesium salts of myoinositol hexakisphosphate is dephosphorylated to provide phosphate and inositol to the growing germ. Dephosphorylation pathways by phosphohydrolases from plants

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and microorganisms have common intermediate metabolites but they also do differ in stereochemistry and in end-products (Greiner et al. 2000, 2002; Quan et al 2003). Greiner et al. (2000) reported that the dephosphorylation is stereochemicallyspecific. Stepwise dephosphorylation of myo-inositol hexakisphosphate by Escherichia coli enzymes yields D/L-Ins(1,2,3,4,5) pentakisphosphate, D/L-Ins(2,3,4,5) tetrakisphosphate, D/L-Ins(2,4,5) trisphosphate or D/L-Ins(1,2,4) trisphosphate, D/LIns(1, 2)-, or (2, 5)-, or (4, 5) bisphosphate, to Ins(2) monophosphate, or Ins(5) monophosphate, where Ins ¼ myo-inositol. Meanwhile, phosphohydrolases from wheat (Triticum aestivum L.) yield D-Ins(1,2,3,5,6) pentakisphosphate, D-Ins (1,2,5,6) tetrakisphosphate, D-Ins(1,2,6) trisphosphate, and D-Ins(1,2) bisphosphate, and Ins(2) monophosphate, whereas the phosphohydrolases LP11 and LP12 from lupin seeds yield D-Ins(1,2,4,5,6) pentakisphosphate, D-Ins(1,2,5,6) tetrakisphosphate, D-Ins(1,2,6) trisphosphate, D-Ins(1,2) bisphosphate, and Ins(2) monophosphate as the final product. Candida phosphohydrolases were observed to yield in sequence: D/L-Ins(1,2,4,5,6) pentakisphosphate, D/L-Ins(1,2,3,5,6) pentakisphosphate, D/L-Ins(1,2,5,6) tetrakisphosphate, D/L-Ins(1,2,5) trisphosphate, D/L-Ins (1, 2) bisphosphate, and D/L-Ins(2) monophosphate (Quan et al. 2003). The various steps of the dephosphorylation of myo-inositol hexakisphosphate are shown as a flow chart to illustrate the sequence that varies in stereochemistry between microorganisms and plant species (Fig. 5.6). In general, enzymes exhibiting an acidic pH optimum release from five to six phosphate groups of myo-inositol hexakisphosphate and myo-inositol monophosphate and myo-inositol. The alkaline phosphohydrolases yield myo-inositol trisphosphate as the end product as these enzymes are unable to use myo-inositol phosphates with three or fewer phosphate groups as substrates (Barrientos et al. 1994).

Ligand Mediation of (or Chemically-Assisted) Enzymatic Dephosphorylation of Complexed Organic Phosphate Monoesters The degradation of organic P-containing substances in the environment is often constrained by physical and biochemical factors affecting the availability of these substrates, more so than any attenuation in the activity of phosphohydrolases and extracellular enzymes in soil (Otani and Ae 1999; Dao 2003, 2004). myo-Inositol hexakisphosphate has six phosphate moieties that provide twelve coordinate ligands for binding metal ions (Fig. 5.7). Metal interactions with the numerous phosphate ligands present on phosphate monoesters can lead to both intra- and intermolecular bonding resulting in the simultaneous formation of numerous monomeric and polymeric species. The phenomenon can lead to the co-precipitation of non-stoichiometric solid-phase mixtures, as the metal to ligand ratio increases. Polyvalent cation binding resulted in reductions in the availability of substrates, shielding organic P from hydrolysis by phosphate monoesterases (Dao 2003, 2007). Phosphate monoesters, having differing number of phosphate moieties, exhibit differing abilities for binding metal ions (Sandberg and Ahderinne 1986;

5 Extracellular Enzymes in Sensing Environmental Nutrients Fig. 5.6 Dephosphorylation pathways of myo-inositol hexakisphosphate by phosphohydrolases in Vicia fava and Lupinus albus. Major pathway denoted by filled solid arrows (Adapted from Greiner et al. (2002))

89

D-Ins(1,2,3,4,5,6)P6 Vicia fava Lupinus alba LP2

Lupinus albus LP11 and LP2

D-Ins(1,2,3,5,6)P5

D-Ins(1,2,4,5,6)P5

D-Ins(1,2,3,6)P4

D-Ins(1,2,5,6)P4

D/L-Ins(1,2,3)P3

D-Ins(1,2,6)P3

D-Ins(1,2)P2

Ins(2)P

Dao 2003), as well as differing affinity for charged surfaces of minerals and soil components (Fig. 5.8) (Anderson et al. 1974; McKercher and Anderson 1989). A consequence of these differences is reflected in their accessibility or susceptibility to enzymatic hydrolysis. In the rhizosphere, plants and microorganisms modify the biochemistry of that environment to affect the solubility and release of complexed organic P and inorganic phosphate. Plant root exudates and siderophores have been long been implicated in the acquisition of P by plants grown in P-deficient soils (Ae et al. 1990; Plessner et al. 1993; Ragothama 1999; Hinsinger 2001; Jones et al. 2003). Excreted organic anions are a variety of products of the citric acid cycle, exchanging with bound phosphate and organic inositol hexakisphosphate anions on soil exchange complex. For example, pigeon pea (Cajanus cajan [L.] Millsp.) releases piscidic acid (p-hydroxybenzyl tartaric acid) and its p-methoxybenzyl derivative in the rhizosphere, allowing the legume to utilize ironbound phosphate. Rape secretes malic and citric acids to release soluble P from a phosphate rock (Hoffland et al. 1989, 1992). Similar organic acids are released by roots of white lupin, when grown on a calcareous P deficient soil (Dinkelaker et al. 1989; Johnson et al. 1996;

90 1.0 0.8 0.6 0.4

PO4 -P relative concentration (C/Co)

Fig. 5.7 Effect of counterions on the dephosphorylation of myoInositol hexakisphosphate by Aspergillus phosphohydrolases (Dao 2003)

T.H. Dao

@ 2:1 Na to IP6-P @ 1:6 Ca to IP6-P @ 3:6 Ca to IP6-P @ 6:6 Ca to IP6-P

0.2 0.0 1.0 @ 2:1 Na to IP6-P @ 1:6 AI to IP6-P @ 3:6 AI to IP6-P @ 6:6 AI to IP6-P

0.8 0.6 0.4 0.2 0.0 1.0 0.8 0.6

@ 2:1 Na to IP6-P @ 1:6 Fe to IP6-P

0.4

@ 3:6 Fe to IP6-P @ 6:6 Fe to IP6-P

0.2 0.0 0

2000

4000

6000

8000

10000

Time (min)

Veneklaas et al. 2003). In soil, commonly identified low molecular weight organic anions include formate, acetate, propionate, oxalate, and citrate, which are formed during microbial metabolism and decomposition of plant residues. Soil organic matter contributes aliphatic acids, phenols, phenolic acids, fulvic and humic substances (Freche et al. 1992; Haynes and Mokolobate 2001). These humic matter ligands are large and have multiple functional groups, binding cations and releasing soluble P to the soil solution. Animal manure and wastewater also contain large amounts of organic matter that likely generate complex ligands (Ma et al. 2001). Owing to the ability of one ligand to participate in exchange reactions with another ligand, polydentate ligands of different size and charge characteristics such as CDTA (1,2-cyclohexane diamino-tetraacetate), DTPA (diethylene triamine-N, N,N0 ,N00 ,N00 -pentaacetate), EDTA (ethylenediamine-N,N,N0 ,N0 -tetraacetate); oxalic acid (ethanedioic acid), and phthalic acid (1,2-benzenedicarboxylic acid) were found to efficiently displace and exchange for myo-inositol hexakisphosphate and other organic phosphates (Dao, 2003, 2004). Adding ligands to a mixture of myo-inositol hexakisphosphate and aluminum reversed the inhibitory effect of Al3þ on the dephosphorylation of the inositol phosphate. When Al3þ concentrations exceeded 0.75 mM, the extent of this phenomenon was in the following order:

5 Extracellular Enzymes in Sensing Environmental Nutrients

a

91

500 400 KH2PO4 Glu-1P α-glycerol-P IP1 IP3 IP6

b

Sorbed concentration (mg P kg–1)

300 200 100 0 500 KH2PO4 Glu-1P α-glycerol-P IP1 IP6

400 300 200 100 0 0

10

20

30

40

50

60

Solution phase P concentration (mg L–1) Fig. 5.8 Sorption of phosphate monoesters, with differing number of phosphate moieties on soil (a) and on peat moss (b) (Adapted from McKercher and Anderson (1989))

phthalate ¼ oxalate < DPTA < EDTA ¼ CDTA. The ligands CDTA and EDTA were able to reverse some, if not most of the inhibition of inositol phosphate hydrolysis at all Al3+ concentrations up to 1.5 mM at pH 4.5 and 6.0. The basis for this reversal of inhibition of dephosphorylation of complexed organic phosphates was found on the stoichiometry of electrical charge. Excess charge was needed to decouple and mobilize complexed myo-inositol hexakisphosphate, and to attain the ligand to myo-inositol hexakisphosphate charge concentration ratios between one and four equivalent molar concentration ratios had to be between 1.5 and 12-fold that of myo-inositol hexakisphosphate, except in the case of phthalate (Fig. 5.9). However, there must be an upper limit to increasing charge concentration needed to overcome the inhibitory effect of polyvalent counterions, because molar concentrations eventually reach levels that would denature or precipitate the enzyme. The reader is referred to a previous review by the author for more detailed information of the chemistry of ligands and the stoichiometry needed to optimize the exchange process (Dao 2007).

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1.0

a

0.8 IP6 alone + CDTA + DTPA + EDTA + Oxalate + Phthalate

0.6 Solution-phase PO4-P relative concentration (C Co –1)

0.4 0.2 0.0 1.0

b

0.8 0.6 0.4 0.2 0.0 1.0

c

0.8 0.6 0.4 0.2 0.0 1.0

d

0.8 0.6 0.4 0.2 0.0 1

2

3

4

Ligand-to-IP6 charge ratio

0

5

10

15

20

25

Equivalent ligand-to-IP6 mole ratio

Fig. 5.9 Effects of selected organic ligands (LIGND), LIGND-to- myo-inositol hexakisphosphate (IP6) charge ratios, and equivalent LIGND-to-IP6 mole ratios on the dephosphorylation of IP6 at four levels of Al3þ counterion, (a ¼ 0; b ¼ 0.25; c ¼ 0.75, and d ¼ 1.5 mM Al3þ), at suspension’s pH 4.5 (Dao 2004)

5.4

An In Vitro Method to Emulate Ligand-Based Plant and Microbial Solubilization and Mineralization of Organic P Pools in Soil and Organic Bio-Nutrient Sources

Although soil enzyme diversity and activities have been used as indicators of ecosystem health (Burns 1982; Dick and Tabatabai 1987; Pankhurst et al. 1997; Dick 1992, 1994; Nannipieri et al. 2002), exogenous enzymes additions have also thought to be quantitative tools to detect changes in organic nutrients in varied environmental media and further elucidate soil nutrient pool sizes, pool turnover rates, and reinforce the role of enzymatic processes in organic nutrient cycling

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(He and Honeycutt 2001; Turner et al. 2002; Dao 2003, 2004; Dao et al. 2006; Green et al. 2006, 2007; Bunemann 2008). In addition, extracellular phosphohydrolases are recently thought of as a means of detecting environmental P accumulation in soil and bioactive forms of P in animal manure and manure-amended soils. Making the concentration of specific enzymes or combinations of enzymes nonlimiting allows quantitative determination of the size of substrate pools beyond chemical identification, as well as allows inference of rates of transformation and transport (Dao 2003, 2004; Dao et al. 2005; Vats et al. 2005; Green et al. 2007; Dao and Hoang 2008). Enzymatic methods have been applied to the identification of organic P-containing substrates in soil extracts, runoff, and surface water columns but have not been used as quantitative methods for soil P pools because of their low efficiency when compared to spectrometric techniques (Pant et al. 1994; He and Honeycutt 2001; Turner et al. 2002; Dao, 2007; Bunemann 2008). For example, Hayes et al. (2000) reported less than 2% of soil total P (6.2 mg kg
1) was desorbed and hydrolyzed in a 50-mM citric acid extract and much less (0.3 mg kg
1) in a water extract. He and Honeycutt (2001) reported a variety of organic P forms in manure and a soil, but achieved a low recovery under their experimental conditions. It must be recognized that the enzymatic intermediates and end products of organic P dephosphorylation are also highly reactive, with many biogeochemical sinks in soils contributing to the low recovery of desorbed and hydrolyzed P. Improvements have recently been made to in situ enzymatic methods by enhancing substrate availability and therefore quantitative recovery of enzyme-labile P content of samples of manure, soils, or runoff (Dao 2003, 2004, 2007; He et al. 2006; Green et al. 2007; Dao and Zhang 2007). Dao (2003) first reported the approach of using polycarboxylate ligands in combination with extracellular enzymes to simultaneously extract, hydrolyze, and categorize P species, without separate steps of an extraction and isolation of the extract for enzymatic assay. The hydrolysis reaction is strongly driven to reach an equilibrium state toward the dephosphorylation of organic P. The rationale and mechanisms for enhanced efficiency have been extensively discussed (Dao 2004, 2007). The in situ experimental approach is distinctly different from other extraction and enzymatic methods (e.g., Hedley et al. 1982; Turner et al. 2002; He et al. 2006). In the two-step methods, the extraction of solid-phase P may not be complete or exhaustive and the extraction process is terminated upon separation and removal of the liquid phase for analysis of extracted P. In this ligand-based fractionation approach, bioavailable phosphate and phosphate monoesters, including inositol phosphates in manure (Dao 2003; Dao et al. 2006) and soils (Dao 2004; Dao et al. 2005; Rao and Dao 2008) can be differentiated into pools that reflect their potential solubilization. Fractions of manure P that are measured in the enzyme-hydrolysable P assay include (1) the water-extractable phosphate, (2) an EDTA-exchangeable phosphate pool that was not previously extracted by water alone, which has been found to be equivalent to the soil P pool extracted by the Mehlich-3 procedure (Dao et al. 2005; Rao and Dao 2008), (3) a water-extractable organic P pool that is hydrolysable by fungal phosphohydrolases, (4) an EDTA-exchangeable organic P pool that is hydrolysable

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by phosphohydrolases and includes inositol phosphates, and (5) a residual pool that is not extractable by water or dilute solutions of polycarboxylate ligands. The ligand-based enzyme-hydrolysable P assay provides a relative scale of chemical and biological stability of P in manures. Phosphorus pools with greater stability, whether the substrates are inorganic phosphates, inositol phosphates, or other organic phosphates, may become biologically available over longer time scales in the soil. This was observed in a case study of the potential for dissolution and solubilization of immobilized P in soils treated with additives to reduce soluble P in soils. A freshly prepared iron hydroxide additive reduced water-extractable P in soil by 90% over a period of 16 weeks. In addition, a plant-available phosphate fraction (Mehlich-3 extractant) was also reduced in iron-treated soils, and both fractions remained unchanged up to 16 weeks. The ligand-based enzyme-hydrolysable P assay, on the other hand, revealed internal changes in P pools and showed that the effect of the iron additive was transitory. The inorganic EDTA extractable P fraction and the enzyme-hydrolysable P pool could be remobilized, reaching initial soil levels to nullify the P-immobilizing action of the iron additives by about the fourth week following soil treatment. The processes of solubilization and desorption of bioavailable P in unamended and Fe(OH)3-amended soils were best described by single or sequential double exponential kinetic equations. This behavior would be consistent with the fact that these substances can range from metastable amorphous to semi crystalline physical states (Arai and Sparks 2002). More evidence of ligand mediation can be found in the enhanced dissolution of minerals (Wieland et al. 1988; Ullman et al. 1996).

5.5

Conclusions

The mineralization of organic matter and organic P compounds, in particular is performed by microorganisms with the ability to hydrolyze phosphorylated organic substrates with the aid of phosphomonoesterase and diesterase enzymes, along with enzymes of many other catabolic processes, occurring simultaneously or in specific sequences. For this reason, the knowledge of functional diversity of the community is as crucial as that of species diversity of that community, the relative contribution of each factor, and finally, the biogeochemical microenvironment they operate in. This chapter reviewed the complementary roles of extracellular enzymes and organic ligands in the solubilization and mineralization of dissolved and insoluble complexes of organic P in agricultural byproducts and in soils and water systems. A high reactivity of organic P with soil constituents and/or suspended colloids resulted in low plant availability and the accumulation of such forms in many soils of the world. Ligand formation, ligand exchange, and extracellular phosphomonoesterase and diesterase enzymes mediate the turnover of organic P. The enzymatic dephosphorylation of uncomplexed organic phosphate monoesters, for example, myo-inositol hexakisphosphate commonly found in soil may yield eventually 6 moles of phosphate and 1 mole of myo-inositol. The metabolites are

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stereochemically-specific, depending on whether the substrate is degraded by microbial systems or whether the substrate is metabolized by plant-derived phosphohydrolases. However, the enzymatic dephosphorylation of organic P-containing substances in the environment often is constrained by physical and biochemical factors affecting the solubility and availability of these substrates. Plant strategies to enhance the sensing and acquisition of P in their roots’ environment have included a wide array of organic ligands and siderophores, and morphological modifications; and there is increasing evidence of complementary genetic adaptation of phosphate sensing and transporter systems, leading to major molecular efforts to research transgenic expression of high-affinity genes induced in response to phosphate deficiency in economically important plants and microbial systems. In furthering our understanding of processes controlling organic P availability, an in situ ligand-based enzymatic method was described, which provides insight into the biological stability of environmental inositol phosphates. Selected polydentate ligands and fungal phosphohydrolases can differentiate various P pools that contribute to the solution-phase phosphate concentration in animal manures and manure-amended soils. A significant amount of intertwining of biotic and abiotic processes was observed in plant and microbes’ arrays of strategies for sensing and obtaining P from their environment. What is clear is that another fundamental evaluation of the biogeochemistry of P in the soil appears necessary to improve our understanding of the solubilization of organic P substrates, and the role of ligands in the turnover of organic P. Although much progress have been made in the past two decades, knowledge gaps in the control mechanisms and the lack of practical effective solutions exist for enhancing the exchange of organic P and their accessibility by extracellular phosphohydrolases, novel, genetically modified, or otherwise. As P fertilizers and supplies are increasingly limited, the challenge to the agricultural communities is to mimic native plant strategies, but on the scale of industrial agricultural production to meet food and fiber needs on the global scale. Soils remain a large pool of stabilized organic P to manage sustainably. This soil P bank is a sizable pool, estimated to average >700 kg ha
1. This pool is relatively stable, or at best, very slowly available. The limited finding the author made on the differential ligand strength between large polycarboxylate and small organic anions such as oxalate, and other aliphatic organic acids (i.e., malate, acetate, etc. . .) in decoupling metal–inositol hexakisphosphate complexes (Dao 2004) apparently indicates a need for new approaches to further enhance the desorption and accessibility to stabilized organic P in soil and sediments. The challenge may include the improvement and translation of our knowledge of surface electrical properties of soil colloids into means and practices to alter the availability of sorbed organic P forms to extracellular enzymes’ catabolic activity. In addition, soil is an extensive repository of a variety of municipal and industrial by-products, i.e., biosolids, coalcombustion ashes, foundry sands; animal manure and wastewater effluents are recycled in production agriculture in large quantities. It is highly desirable is highly desirable to continue research on the effect of soil management and extracellular enzymes of the major nutrient cycles of C and N, in addition to phosphohydrolases. It is imperative we improve our holistic understanding of nutrients’ interaction on

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soil enzymes’ induction and release in order to find solutions to constraints to their activity and the cycling of P to improve the use efficiency of this non-renewable resource.

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Tadano T, Sakai H (1991) Secretion of acid phosphatases by roots of several crop species under phosphorus deficient conditions. Soil Sci Plant Nutr 37:129–140 Tanaka K, Yoshida T, Kasai Z (1974) Radioautographic demonstration of the accumulation site of phytic acid in rice and wheat grains. Plant Cell Physiol 15:147–151 Tarafdar JC, Claassen N (2003) Organic phosphorus utilization by wheat plants under sterile conditions. Biol Fertil Soils 39:25–29 Tarafdar JC, Marschner H (1994) Phosphatase activity in the rhizosphere and hyphosphere of VA mycorrhizal wheat supplied with inorganic and organic phosphorus. Soil Biol Biochem 26:381–395 Ternan NG, Mc Grath JW, Mc Mullan G, Quinn JP (1998) Review: organo-phosphonates: occurrence, synthesis and biodegradation by microorganisms. World J Microbiol Biotechnol 14:635–647 Torrent J, Barren V, Schwertmann U (1990) Phosphate adsorption and desorption by goethites differing in crystal morphology. Soil Sci Soc Am J 54:1007–1012 Turner BL, McKelvie ID, Haygarth PM (2002) Characterization of water-extractable soil organic phosphorus by phosphatase hydrolysis. Soil Biol Biochem 34:27–35 Turner BL, Mahieu N, Condron LM (2003) Phosphorus-31 nuclear magnetic resonance spectral assignments of phosphorus compounds in soil NaOH–EDTA extracts. Soil Sci Soc Am J 67:497–510 Turner BL, Frossard E, Baldwin DS (2005) Organic phosphorus in the environment. CABI, Oxfordshire, UK Turner BL, Richardson AE, Mullaney EJ (2007) Inositol phosphates: linking agriculture and environment. CABI, Oxfordshire, UK Ullman WJ, Kirchman DL, Welch SA, Vandevivere P (1996) Laboratory evidence for microbially mediated silicate mineral dissolution in nature. Chem Geol 132:11–17 Vats P, Bhattacharyya MS, Banerjee UC (2005) Use of phytases (myo-inositol hexakisphosphate phosphohydrolases) for combating environmental pollution: a biological approach. Crit Rev Environ Sci Technol 35:469–486 Veneklaas EJ, Stevens J, Cawthray GR, Turner S, Grigg AM, Lambers H (2003) Chick pea and white lupin rhizosphere carboxylates vary with soil properties and enhance phosphorus uptake. Plant Soil 248:187–197 Voglmaier SM, Bembenek ME, Kaplin AI, Dorman G, Olszewski JD, Prestwich GD, Snyder SH (1996) Purified inositol hexakisphosphate kinase is an ATP synthase: diphosphoinositol pentakisphosphate as a high-energy phosphate donor. Proc Natl Acad Sci 93:4305–4310 Vohra A, Satyanarayana T (2003) Phytases: microbial sources, production, purification, and potential biotechnological applications. Crit Rev Biotechnol 23:29–60 Wasaki J, Yamamura T, Shinano T, Osaki M (2003) Secreted acid phosphatase is expressed in cluster roots of lupin in response to phosphorus deficiency. Plant Soil 248:129–136 Weaver JD, Mullaney EJ, Lei XG (2007) Altering the substrate specificity site of Aspergillus niger PhyB shifts the pH optimum to pH 3.2. Appl Microbiol Biotechnol 76:117–122 White AK, Metcalfe WW (2007) Reduced phosphorus compounds. Annu Rev Microbiol 61:379–400 Wieland E, Wehrli B, Stumm W (1988) The coordination chemistry of weathering. III. A generalization on the dissolution rates of minerals. Geochim Cosmochim Acta 52:1969–1981 Wild A, Oake OL (1966) Organic phosphate compounds in calcium chloride extracts of soils: identification and availability to plants. J Soil Sci 17:356–371 Yamagata H, Tanaka K, Kasai Z (1980) Purification and characterization of acid phosphatase in aleurone particles of rice grains. Plant Cell 21:1449–1460 Zhang H, Dao TH, Basta NT, Dayton EA, Daniel TC (2006) Remediation techniques for manure nutrient loaded soils. In: Rice JM, Cadwell DF, Humenik FJ (eds) National Center for Manure and Animal Waste Management White Papers. American Society Agriculture Biological Engineers. Pub. 913C0306. St. Joseph, MI, pp 483–503

Chapter 6

Importance of Extracellular Enzymes for Biogeochemical Processes in Temporary River Sediments during Fluctuating Dry–Wet Conditions Annamaria Zoppini and J€ urgen Marxsen

6.1

Introduction

The special hydrological circumstances that characterize temporary rivers (Fig. 6.1) make them particularly sensitive to environmental alterations. Temporary waters are found throughout the world and include intermittent streams and ponds, episodic rain puddles and seasonal limestone lakes. These natural bodies of water experience a recurrent dry phase of varying duration and spatial extent (Uys and O’Keeffe 1997). Temporary waters are widespread in semi-arid regions worldwide where they play an important role as water sources (e.g., water supply, irrigation and hydroelectric power generation). Climate change represents an emerging problem for these ecosystems by increasing the frequency and duration of drought periods, with potentially important effects on fluvial biogeochemical processes. From the examination of new findings from the past 6 years of research the Intergovernmental Panel on Climate Change Working Group I (IPCC WGI 2007) concluded that warming of the climate system is unequivocal. An ensemble of 12 climate models (Milly et al. 2005) projects changes in stream flow with 10–30% decreases in runoff in southern Europe by the year 2050. This trend has already been experienced over the last century not only in the Mediterranean basins (WFD/ EUWI 2006), but also in streams and rivers in temperate European regions. Here decreasing runoffs and extended periods of desiccation have been observed, especially in headwaters (Wilby et al. 2006; Sutherland et al. 2008).

A. Zoppini (*) Istituto di Ricerca Sulle Acque, Consiglio Nazionale delle Ricerche, Area della Ricerca di Roma 1, via Salaria Km 29.300, CP10, 00015 Monterotondo, Roma, Italy e-mail: [email protected] J. Marxsen Limnologische Fluss-Station des Max-Planck-Instituts f€ ur Limnologie, 36110 Schlitz, Germany Institut f€ur Allgemeine und Spezielle Zoologie, Tier€ okologie, Justus-Liebig-Universit€at Gießen, Heinrich-Buff-Ring 26–32, 35392 Gießen, Germany

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Fig. 6.1 Changes in hydrological conditions in temporary rivers (after Kirkby and Froebrich (2006), modified)

Temporary rivers are the dominant surface water bodies in the Mediterranean region although they are rarely monitored. Until recently dry reaches have been described as “biologically inactive” (Stanley et al. 1997) and there is still little information available on the biogeochemical processes. Tzoraky et al. (2007) observed that in semi-arid catchments the channel bed processes continue even at a low level (40%) of sediment moisture. As a result, the first water flush entering into contact with sediments after a drought period could be extremely modified in its chemistry compared to the base flow, causing in turn drastic modifications in the chemistry of receiving water-bodies (e.g., lakes, rivers, and coastal waters) (Fig. 6.1). Aquatic sediments act as a sink and source of nutrients. Microbial degradation and transformation of the organic matter deposited in a river channel bed are key processes with regard to the carbon cycling in the lotic food web, which links sedimentary organic matter to the upper level of the community, including carnivores (Fischer and Pusch 2001; Findlay et al. 2003; Mulholland 2003; Marxsen 2006). Via the microbial food chain, complex organic substrates are solubilized by extracellular enzymes in a series of steps from particulate organic matter to high molecular weight dissolved organic carbon and low molecular weight substrates (Chrost 1991). A strong coupling between bacterial activities and deposition of organic material in sediment has been found for both marine and freshwater systems (Sander and Kalff 1993; Goedkoop et al. 1997; Wobus et al. 2003). Hence the efficiency of extracellular enzymes represents a critical step and is able to influence the incorporation of organic carbon into bacterial cells and the consequent transfer to the food chain (Marxsen and Fiebig 1993). Drying and rewetting are well known climatic factors influencing the physiological status of microbial biomass in soil ecosystems, with significant effects on mineralization (e.g., Raubuch et al. 2002; Griffiths et al. 2003; Mikha et al. 2005). Although benthic microbial processes have been recognized as having an important role in carbon and nutrient flux, few studies have documented the

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response to dry–wet cycles in temporary waters. The exact role of microbes in mediating these processes is still largely unresolved because methodological constraints make it difficult to determine whether the effects are biologically or physically driven. In addition, in situ measurements are very difficult because of high temporal variability and spatial heterogeneity. This issue has been recently addressed by the European Commission through its funding of Research Projects on temporary rivers (TempQsim and Mirage) with the aim to provide advanced tools to significantly improve the efficiency of integrated water management in Mediterranean semiarid river catchments. This action has encouraged experimental studies on the role of the benthic microbial community in carbon, nutrient and energy flux during drastic changes in water availability (Tzoraky et al. 2007; Amalfitano et al. 2008; Fazi et al. 2008; Zoppini et al. 2010).

6.2

The Response of Benthic Microbial Communities to Drought: A Matter of Survival

Drought has been cited as the most serious of natural disasters as regards loss of life and its impact on agricultural production and economics (Wilhite 2000), although it is a natural feature of aquatic ecosystems in most regions of the world (McMahon and Finlayson 2003). Temporary waters result in sediment desiccation during an extended period of the year with the consequence of exposing the benthic organisms living there to the air. Biota that inhabits these ecosystems must be morphologically, physiologically and behaviorally adaptable to survive such conditions until the first flood arrives. Desiccation has been reported to alter the chemistry (e.g., De Groot and Van Wijck 1993) and mineralogy (e.g., Baldwin 1996) of the sediment or soil and kills up to three-quarters of the microbes (e.g., Qiu and McComb 1995). In semiarid regions of the planet drought events have a major effect on benthic community functions, including a delay in litter decomposition together with a decrease in invertebrate density (Pinna and Basset 2004; Fonnesu et al. 2004; Larned et al. 2007). Tolerance mechanisms to desiccation are poorly understood despite the fact that numerous prokaryotic and eukaryotic organisms are capable of surviving more or less complete dehydration. Drying-rewetting imposes physiological constraints that few genera of bacteria, called anhydrobiotes, can tolerate (Potts 1994; Billi and Potts 2002). Bacteria can cope with this problem through strategies such as forming polymers and spores with the ability to resist physical blows. The production and storage of intracellular solutes, like amino acids and low molecular weight carbohydrates, for acclimatizing to low water conditions (Halverson et al. 2000), is especially important for Gram-negative bacteria, which are unable to form spores and are more susceptible than Gram-positives to disruption by osmotic stress because of their less stable cell walls (Fierer et al. 2003; Schimel et al. 2007).

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Examples of spore-forming bacteria are quite widespread among Gram-positive bacteria and they colonize various habitats, including the aquatic environment. In their dormant state, spores have no detectable metabolism although recent findings indicate that dormant bacterial spores, belonging to the genus Bacillus, can significantly influence the distribution of heavy metals in the sedimentary marine environment by enzymatic catalysis (Francis and Tebo 2002; Dick et al. 2006). Moreover over the long term (millions of years) catalase was found in freezedried permafrost samples (Gilichinsky et al. 1992), while in deep-sea sediments enzyme activity was detected even in a 124,000 year-old sapropel layer (Coolen and Overmann 2000). Overall, these studies suggest that the commonly held view that bacteria in the dormant state are inactive should be revised. Lowland river–floodplain systems are characterized by a high degree of variability where inundation marks the shift from a terrestrial ecosystem to an aquatic one. In these systems partial drying of wet (previously inundated) sediments results in an increased sediment affinity for phosphorus and produces a zone where there is nitrification coupled with denitrification causing a reduction of nutrient availability (Baldwin and Mitchell 2000). A complete desiccation of sediments may lead to the death of bacteria (and subsequent mineralization of nutrients), a decrease in the affinity of P for iron minerals, a decrease in microbial activity and a cessation of all anaerobic bacterial processes (e.g., denitrification). For semi-permanent stream sediments Rees et al. (2006) showed that changes in microbial community structure were preserved even one month after rewetting, with a significant difference from the pre-drought and drought microbial communities. The functional properties of microbial communities were investigated in the sediment of ephemeral rivers (Larned et al. 2007). Non-specific esterase activity, used as an assay for total enzyme activity, was negatively related to the dry period length. The esterase activity decreased by a negative exponential model in the dry periods of 1–7 days accompanied by decreasing respiration rates. Moreover a prolonged period of drought (up to 417-days) further affected esterase activity, reducing the rates by an order of magnitude to close to the detection limit. Extracellular enzymes may be stable in dry soils or aquatic biofilms for weeks. For phosphatases Perez-Mateos et al. (1991) observed that 65% of indigenous enzymes remained active after 50 days of soil storage at 22
C, while Romanı´ and Sabater (1997) found that extracellular enzyme activity recovered immediately in stromatolitic riverine communities when rewetted after a summer drought. Benthic cyanobacterial mats from marshes of northern Belize, an oligotrophic environment, are exposed to extreme conditions in terms of hydrology, nutrient availability and salinity (Sirova´ et al. 2006). In the cyanobacterial mats alkaline phosphatase exhibited the highest extracellular enzyme activity, followed by leucineaminopeptidase, arylsulphatase, and b-glucosidase. During the period of drought dry mats retained the same level of potential phosphatase activity after 10 weeks of desiccation (350–450 mmol MUF g1 ash free dry weight min1). In this environment the preservation of enzyme activity may be favored by the levels of polysaccharide-rich extracellular polymeric substances (EPS) that characterise benthic

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cyanobacterial mats. Most of the phosphatase activity, visualized using artificial ELF®97 phosphate, appeared free and located in the EPS matrix throughout the mat, with a decoupling from its source in both space and time. Many studies on the effect of droughts on benthic microorganisms encounter difficulties in disentangling the relative effects of the spatial and temporal extents of low flows and of these from the river bed drying pattern. The alternative approach to gathering information is to simulate a drought event in the laboratory. Amalfitano et al. (2008) collected wet sediments from four temporary European rivers and let them dry under controlled conditions until complete desiccation. Despite the different origin of the microbial communities the responses of structural and functional parameters to drought were very similar (Fig. 6.2). Bacterial carbon production exponentially decreased, nearly ceasing in dry conditions, followed by a slower decrease in bacterial abundance, with an overall reduction of 74%. By the end of the experiment, live cells (14% of the initial value) were depressed in their main metabolic functions. Hence a conspicuous number of live cells were still abundant at the end of the experiment, but mostly depressed in their metabolic activity. As a result, the significant decrease in per-cell production resulted in a very substantial increase in the community turnover time. Community composition shifted, with an increase in Alpha- and Betaproteobacteria when sediment was dried. In the same experiment extracellular enzyme activities, measured fluorometrically (Wobus et al. 2003), involved in P, N, and C cycling were weakly or not at all affected by the progressive decreasing of water availability (Zoppini 2007). Aminopeptidase activity was affected by drying and displayed a slow decreasing trend in all tested rivers although it preserved between 40 and 60% of the initial rates in dry sediments (Fig. 6.3). Overall the capacity of alkaline phosphatase to hydrolyze phosphorilated organic matter did not change significantly during the progressive loss of moisture: only one in four tested sediments (the river Krathis) showed a negative trend with desiccation but still preserved 80% of its initial capacity during dry conditions. Similarly the hydrolyzing capacity of polymers like polysaccharides and lipids (b-glucosidase and lipase activities) was preserved under drought conditions without showing any significant trend. This enzyme activity survival capacity recalls mechanisms found in desiccation-tolerant bacterial cells capable of accumulating proteins, of which many are able to remain stable (Billi and Potts 2002), or the capacity of polysaccharides-rich extracellular polymeric substances (EPS) to favor enzyme preservation (Sirova´ et al. 2006). To sum up the data, microbial communities are significantly affected by water stress conditions and there are changes in microbial community structure with a drastic reduction of cell abundance, vitality and metabolic activity. Hydrolytic enzymes constitute an exception to this trend. They are probably preserved even if the cells in which they originated become non-viable. We postulate that the preservation of these enzymes represents an important mechanism for the fast recovery of surviving microbial cells after drought. If the hydrolysis of organic matter continues in the dry sediment we can infer that the pool of organic compounds contained in the first water flush could be enriched with labile nutrients that accumulate in the sediments as bacteria are died or temporarily

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Fig. 6.2 Total and live (Live/Dead® BacLight™ dye technique) bacterial cell abundance (top panel) and carbon production (3H-Leucine inc.) (bottom panel) versus sediment water content, expressed as a percentage of sediment water hold capacity (WHC). Note the x-axis reverse scale: 100% identifies wet sediment at the beginning of the experiment, 0% corresponds to the ending dry sediment. All data are normalized to grams of dry sediment. Error bars indicate standard deviations of three independent measurements. Regression curves are shown originating from temporary rivers (total cell abundance: r2 ¼ 0.69, P < 0.05; live cell abundance r2 ¼ 0.76, P < 0.05; 3Hleucine inc. r2 ¼ 0.79, P < 0.05). (square Tagliamento, Italy; triangle Krathis, Greece; diamond Mulargia, Italy; circle Pardiela, Portugal). (after Amalfitano et al. (2008), modified)

inactive. However, in the case of a complete stop of extracellular enzyme activity during extreme dryness, enzymes are able to become active immediately upon rewetting (Marxsen et al. 2010), thus resulting in immediate delivery of these compounds.

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Fig. 6.3 Extracellular enzyme activities versus sediment water content expressed as percentage of sediment water hold capacity (WHC) (see Fig. 3 for symbols). All data are normalized to gram of

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Benthic Microbial Community Awakening After Flooding

The intense run-off and flushing associated with the heavy storms typically occur shortly after the end of a dry period. These events can severely impact microbial distribution as river discharges can increase by several orders of magnitude with respect to their regular flow (Holmes et al. 1998). Inundation of previously dry river reaches can trigger ecological “hot moments” during which biogeochemical reactions or biological processes begin to accelerate after long quiescent periods (McClain et al. 2003). Inundation can activate microbial and algal cells, delivers chemical substrates to reaction sites, and stimulates enzymecontrolled nutrient transformations and organic matter mineralization (Baldwin and Mitchell 2000; Burns and Ryder 2001; Belnap et al. 2005; Romanı` et al. 2006). Several findings suggest that biota response to inundation depends on the duration of the preceding dry phase (Baldwin and Mitchell 2000; Larned et al. 2007). After 17 days of inundation esterase enzyme activity in sediments that had been dry for 256–523 days was comparable to the potential activity in non-inundated sediments from a similar dry-period range (Larned et al. 2007). After the same dry-period range and 17 days of inundation there was no detectable relationship between sediment respiration rate and dry period duration, although rates tended to be higher for inundated sediments than for non-inundated sediments. Strong temporal peaks in enzyme activity were observed in sediments within 7 days following inundation (Burns and Ryder 2001). The short response time of a- and b-glucosidase activities after 24 h flooding suggests that even short pulses in high flows may stimulate bacterial activity as dissolved organic carbon loads also peak at this time. However, a longer wetting time may be needed to drive hydrolysis of the proteins, fatty acids and longer chain polysaccharides. This indicates a rapid use of available carbon by microbial communities. A general decline in enzyme activity rates was found in the 21 days following the first week from inundation, which was thought to be the result of substrate limitation within the flooded cores or inhibition by hydrolysis end products. There have been laboratory-scale experiments conducted on dry sediment simulating an inundation event in order to demonstrate the role of sediments as a source of microbial populations, and related activities, in “first-flush” water (Fazi et al. 2008). Within 9 h after inundation of dry sediments, benthic bacteria colonized the overlaying water and approximately 20% of total cells exhibited DNA de novo synthesis (bromodeoxyuridine-positive). The primary microbial colonizers of the overlaying water – as determined by 16S rRNA gene sequence analysis – were related to at least six different phylogenetic lineages of Bacilli, a group with many

Fig. 6.3 (continued) dry sediment. Data reported are from duplicated microcosms and single values are means of four measurements. Regression curves are shown (aminopeptidase r2 ¼ 0.78 Pardiela river and r2 ¼ 0.52 in the rest of rivers, P < 0.05; alkaline phosphatase r2 ¼ 0.80, P < 0.05) (Zoppini 2007)

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spore-forming members (Onyenwoke et al. 2004), and Alphaproteobacteria (Brevundimonas spp. and Caulobacter spp.). The microbial awakening was accompanied by C production rates similar to those measured in highly productive eutrophic systems with a prevalence of biomass synthesis (3H-leucine incorporation) over cell division (3H-thymidine incorporation) (Fig. 6.4). Significant extracellular enzyme activities were also observed: within 28 h after inundation aminopeptidase activity reached 70% of the peak value, which was after 72 h (431 nmol MCA L1 h1), while alkaline phosphatase activity reached 83% (99 nmol MUF l1 h1). The analysis of the aminopeptidase to alkaline phosphatase activity ratio showed the different role played by these enzymes in the metabolic awakening of bacterial cells. The lower hydrolysis rate of organic phosphorus compared to proteins, described by the increasing AMA:APA ratio, indicated a slower P-remobilization compared to N-remobilization. Aminopeptidase and alkaline phosphatase activities were also significantly correlated to bacterial carbon and cell production (P < 0.01; n ¼ 12). This confirmed the key role played by extracellular enzymes in making available organic compounds for both the synthesis of intracellular proteins and the production of new biomass. A more advanced tool for studying metabolic properties in streambed sediments is represented by the perfused core technique (Marxsen and Fiebig 1993). The application of this approach enables the acquisition of new information on the time and mode of recovery of benthic microbial communities after flooding without major disturbances of the sediment core (Marxsen et al. 2010). The response of two river sediments, originating from semi-arid (Mulargia, Italy) and temperate (Breitenbach, Germany) climatic regions, were analyzed. Both sediments were similar in microbial community composition, determined via CARD-FISH, in that they were dominated by Betaproteobacteria (39–45%) and Alphaproteobacteria (27–31%). However, the Mulargia sediment contained a higher percentage of Gram-positive bacteria (24%) than the Breitenbach one (9%). After rewetting bacterial cell abundances did not change significantly from their initial values, which were similar to those observed in the wet sediments (Mulargia 10  108 cells mL1 sediment and Breitenbach 30  108 cells mL1 sediment) (Fig. 6.5). The functional awakening of the microbial community was marked by rapid bacterial carbon production, which reached the maxima rates within 48 h in both sediments (Fig. 6.5). This trend was also accompanied by a rapid increase in extracellular enzyme activities (Fig. 6.6). A few hours after flooding aminopeptidase activity reached about half the level measured in non-desiccated sediments. This activity increased further until the end of the experiment, when values reached those measured in unaffected sediments. Alkaline phosphatase was reactivated within a few hours although in both sediments it underwent a progressive decrease. These experimental findings are in accordance with previous observations. Alkaline phosphatase is involved in phosphorous remobilization and its activity is proportional to phosphorous demand. It has been estimated that 30–60% of the microbial biomass carbon contained in the soil may be released during an individual rewetting event along with water soluble phosphorous deriving, for example, from the

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Fig. 6.4 (a) Extracellular enzyme activities (AMA, aminopeptidase activity; APA, alkaline phosphatase activity). (b) Bacterial carbon production (3H-leucine inc.) and cell production (3Hthymidine inc.). (c) The AMA to APA ratios and 3H-Leu to 3H-TdR ratios. Data are means and standard deviation of duplicate measurements from three independent microcosms (after Fazi et al. (2008), modified)

microbial cell rupture caused by osmotic shock (Kieft et al. 1987; Halverson et al. 2000; Baldwin and Mitchell 2000; Austin et al. 2004). The hydrolysing activity related to polysaccharides (b-glucosidase) and lipids (lipase) also showed a fast recovery with differences among sediments (Fig. 6.6). The high activity levels measured at the beginning of rewetting in both

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Bacterial abundance (108 cells µL–1)

100 Breitenbach M

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Bacterial carbon production (µgCh–1 µL–1)

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2.52

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1.0 Breitenbach

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Fig. 6.5 (a) Bacterial abundance and (b) bacterial carbon production (BCP, 14C-leucine incorporation) after rewetting of desiccated streambed sediments. Average values and standard deviations are shown (if not visible, sd is hidden behind symbols). The columns give data from non-desiccated sediments (B Breitenbach, M Mulargia) (after Marxsen et al. (2010), modified)

environments, especially those for enzymes involved in polymer degradation (bglucosidase, peptidase and lipase), can be taken as an indication of outlasting of extracellular enzymes during drought, confirming previous findings on the preservation of enzymes during drying (Sirova´ et al. 2006; Fazi et al. 2008; Zoppini 2007). Their activities can play an important role in fuelling the bacterial metabolism as they supply organic substrates rich in energy (glucose and fatty acids) and amino acids ready to use for synthesising new biomass.

6.4

Conclusions

Overall our understanding of the effect of dry–wet conditions on the ecology of temporary rivers is still limited. From the information available on the functional properties of microbial communities we can infer that their role in carbon, nutrient and energy flux in water stress conditions is important.

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aminopeptidase (nmol AMC h–1 µL–1)

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Fig. 6.6 Extracellular enzyme activities after rewetting of desiccated streambed sediments. Average values and standard deviations are shown (if not visible, sd is hidden behind symbols). The columns give data from non-desiccated sediments (B Breitenbach, M Mulargia) (after Marxsen et al. (2010), modified)

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A common feature of rewetted sediments (and soil) is a large flush of mineral phosphorous and nitrogen in the initial phase. Microbial communities can significantly contribute to this via different mechanisms. The preservation of enzymes during periods of drought, even in prohibitive conditions for the rest of the microbial metabolism, determines an excess of labile organic matter and nutrients ready to be taken up later by bacteria at water arrival or to be delivered immediately upon rewetting. In this scenario the first flood delivers labile low molecular organic matter and nutrients to the receiving water bodies with the potential to accelerate microbial processes and affect water quality (i.e., the onset of anoxia). The fast resumption of enzyme activity favors the rapid re-establishment of sediment functions due to aquatic microbial communities. It appears that dryingrewetting plays a key role in a number of metabolic processes in temporary rivers and the length of drought periods is important in affecting biogeochemical cycling. Increased periods of drought, due to climate change, could thus represent a bottleneck for bacterial communities living in temporary waters and they will need to adopt their metabolic strategies to survive them.

References Amalfitano S, Fazi S, Zoppini A, Barra Caracciolo A, Grenni P, Puddu A (2008) Responses of benthic bacteria to experimental drying in sediments from Mediterranean temporary rivers. Microb Ecol 55:270–279 Austin AT, Yahdjian L, Stark JM, Belnap J, Porporato A, Norton U, Ravetta DA, Schaeffer SM (2004) Water pulses and biogeochemical cycles in arid and semiarid ecosystems. Oecologia 141:221–235 Baldwin DS (1996) Effects of exposure to air and subsequent drying on the phosphate sorption characteristics of sediments from a eutrophic reservoir. Limnol Oceanogr 41:1725–1732 Baldwin DS, Mitchell AM (2000) The effects of drying and re-flooding on sediment and soil nutrient dynamics of lowland river–floodplain systems: a synthesis. Regulated Rivers: Res Manag 16:457–467 Belnap J, Welter JR, Grimm NB, Barger N, Ludwig JA (2005) Linkages between microbial and hydrologic processes in arid and semiarid watersheds. Ecology 86:298–307 Billi D, Potts M (2002) Life and death of dried prokaryotes. Res Microbiol 153:7–12 Burns A, Ryder DS (2001) Response of bacterial extracellular enzymes to inundation of floodplain sediments. Freshw Biol 46:1299–1307 Chrost RJ (1991) Environmental control of the synthesis and activity of aquatic microbial ectoenzymes. In: Chrost RJ (ed) Microbial enzymes in aquatic environments. Springer, New York, pp 29–59 Coolen MJL, Overmann J (2000) Functional exoenzymes as indicators of metabolically active bacteria in 124,000-year-old sapropel layers of the eastern Mediterranean sea. Appl Environ Microbiol 66:2589–2598 De Groot C, Van Wijck C (1993) The impact of desiccation of a freshwater marsh (Garcines Nord, Camargue, France) on sediment–water–vegetation interactions. Part one: the sediment chemistry. Hydrobiologia 252:83–94 Dick GJ, Lee YE, Tebo BM (2006) Manganese(II)-oxidizing bacillus spores in Guaymas Basin hydrothermal sediments and plumes. Appl Environ Microbiol 72:3184–3190 Fazi S, Amalfitano S, Piccini C, Zoppini A, Puddu A, Pernthaler J (2008) Colonization of overlaying water by bacteria from dry river sediments. Environ Microbiol 10:2760–2772

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Fierer N, Schimel JP, Holden PA (2003) Influence of drying-rewetting frequency on soil bacterial community structure. Microb Ecol 45:63–71 Findlay SEG, Sinsabaugh RL, Sobczak WV, Hoostal M (2003) Metabolic and structural response of hyporheic microbial communities to variations in supply of dissolved organic matter. Limnol Oceanogr 48:1608–1617 Fischer H, Pusch M (2001) Comparison of bacterial production in sediments, epiphyton and the pelagic zone of a lowland river. Freshw Biol 46:1335–1348 Fonnesu A, Pinna M, Basset A (2004) Spatial and temporal variations of Detritus breakdown rates in the river Flumendosa Basin (Sardinia, Italy). Int Rev Hydrobiol 89:443–452 Francis CA, Tebo BM (2002) Enzymatic manganese(II) oxidation by metabolically dormant spores of diverse Bacillus species. Appl Environ Microbiol 68:874–880 Gilichinsky DA, Vorobyova EA, Erokhina LG, Fyodorov-Dayvdov DG, Chaikovskaya NR (1992) Long-term preservation of microbial ecosystems in permafrost. Adv Space Res 12:255–263 Goedkoop W, Gullberg KR, Johnson RK, Ahlgren I (1997) Microbial response of a freshwater benthic community to a imulated diatom sedimentation event: interactive ejects of benthic fauna. Microb Ecol 34:131–143 Griffiths RI, Whiteley AS, O’Donnell AG, Bailey MJ (2003) Physiological and community responses of established grassland bacterial populations to water stress. Appl Environ Microbiol 69:6961–6968 Halverson LJ, Jones TM, Firestone MK (2000) Release of intracellular solutes by four soil bacteria exposed to dilution stress. Soil Sci Soc Am J 64:1630–1637 Holmes RM, Fisher SG, Grimm NB, Harper BJ (1998) The impact of flash floods on microbial distribution and biogeochemistry in the parafluvial zone of a desert stream. Freshw Biol 40:641–654 IPCC (2007) Climate change 2007: the physical science basis. Contribution of working group I to the fourth assessment report of the intergovernmental panel on climate change. Cambridge University Press, Cambridge, 996 Kieft T, Soroker E, Firestone M (1987) Microbial biomass response to a rapid increase in water potential when dry soil is wetted. Soil Biol Biochem 19:119–126 Kirkby M, Froebrich J (2006) Introduction. In: Froebrich J, Bauer M (eds) Critical issues in the water quality dynamics of temporary waters – evaluation and recommendations from the TempQsim Project. Booklet – enduser summary. Institute for water quality and waste management, University of Hannover, Hannover, pp 8–11 Larned ST, Datry T, Robinson C (2007) Invertebrate and microbial responses to inundation in an ephemeral river reach in New Zealand: effects of preceding dry periods. Aquat Sci 69:554–567 Marxsen J (2006) Bacterial production in the carbon flow of a central European stream, the Breitenbach. Freshw Biol 51:1838–1861 Marxsen J, Fiebig DM (1993) Use of perfused cores for evaluating extracellular enzyme activity in stream-bed sediments. FEMS Microbiol Ecol 13:1–11 Marxsen J, Zoppini A, Wilczek S (2010) Microbial communities in streambed sediments recovering from desiccation. FEMS Microb Ecol 71:374–386 McClain ME, Boyer EW, Dent CL, Gergel SE, Grimm NB, Groffman PM, Hart SC, Harvey J, Johnston CA, Mayorga E, McDowell WH, Pinay G (2003) Biogeochemical hot spots and hot moments at the interface of terrestrial and aquatic ecosystems. Ecosystems 6:301–312 McMahon TA, Finlayson BL (2003) Droughts and anti-droughts: the low-flow hydrology of Australian rivers. Freshw Biol 48:1147–1160 Mikha MM, Riceb CW, Millikenc GA (2005) Carbon and nitrogen mineralization as affected by drying and wetting cycles. Soil Biol Biochem 37:339–347 Milly PCD, Dunne KA, Vecchia AV (2005) Global pattern of trends in streamflow and water availability in a changing climate. Nature 438:347–350 Mulholland PJ (2003) Large-scale patterns in dissolved organic carbon concentration, flux and sources. In: Findlay SE, Sinsabaugh RL (eds) Aquatic ecosystems, Interactivity of dissolved organic matter. Academic, Elsevier, San Diego, pp 139–159

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Onyenwoke RU, Brill JA, Farahi K, Wiegel J (2004) Sporulation genes in members of the low G+C Gram-type- positive phylogenetic branch (Firmicutes). Arch Microbiol 182:182–192 Perez-Mateos M, Busto MD, Rad JC (1991) Stability and properties of alkaline phosphate immobilized by a rendzina soil. J Sci Food Agr 55:229–240 Pinna M, Basset A (2004) Summer drought disturbance on plant detritus decomposition processes in three River Tirso (Sardinia, Italy) sub-basins. Hydrobiologia 522:311–319 Potts M (1994) Desiccation tolerance of prokaryotes. Microbial Rev 58:755–805 Qiu S, McComb AJ (1995) Planktonic and microbial contributions to phosphorus release from fresh and air-dried sediments. Mar Freshw Res 46:1039–1045 Raubuch M, Dyckmans J, Joergensen RG, Kreutzfeldt M (2002) Relation between respiration, ATP content, and adenilate energy charge (AEC) after incubation at different temperature and after drying and rewetting. J Plant Nutr Soil Sci 165:435–440 Rees GN, Watson GO, Baldwin DS, Mitchell AM (2006) Variability in sediment microbial communities in a semipermanent stream: impact of drought. J North Am Benthol Soc 25:370–378 Romanı` AM, Sabater S (1997) Metabolism recovery of a stromatolitic biofilm after drought in a Mediterranean stream. Archiv f€ ur Hydrobiologie 140:262–271 Romanı` AM, Vazquez E, Butturini A (2006) Microbial availability and size fractionation of dissolved organic carbon after drought in an intermittent stream: biogeochemical link across the stream–riparian interface. Microb Ecol 52:501–512 Sander BC, Kalff J (1993) Factors controlling bacterial production in marine and freshwater sediments. Microb Ecol 26:79–99 Schimel J, Balser TC, Wallenstein M (2007) Microbial stress–response physiology and its implications for ecosystem function. Ecology 8:1386–1394 Sirova´ D, Vrba J, Rejma´nkova´ E (2006) Extracellular enzyme activities in benthic cyanobacterial mats: comparison between nutrient-enriched and control sites in marshes of northern Belize. Aquat Microb Ecol 44:11–20 Stanley EH, Fisher SG, Grimm NB (1997) Ecosystem expansion and contraction in streams. Bioscience 47:427–435 Sutherland WJ, Bailey MJ, Bainbridge IP et al (2008) Future novel threats and opportunities facing UK biodiversity identified by horizon scanning. J Appl Ecol 45:821–833 Tzoraky O, Nikolaidis NP, Amaxidis Y, Skoulikidis NTH (2007) In-stream biogeochemical processes of a temporary river. Environ Sci Technol 41:1225–1231 Uys CM, O’Keeffe J (1997) Simple words and fuzzy zones: early directions for temporary river research in South Africa. Environ Manage 21:517–531 WFD/EUWI (2006) Mediterranean joint process WFD/EUWI, Water Scarcity Drafting Group, Tool Box (Best practices) on water scarcity. Draft Version 9, to be modified, 13 Feb 2006 Wilby RL, Whitehead PG, Wade AJ, Butterfield D, Davis RJ, Watts G (2006) Integrated modelling of climate change impacts on water resources and quality in a lowland catchment: River Kennet, UK. J Hydrol 330:204–220 Wilhite DA (2000) Drought as a natural hazard. Concept and definition. In: Wilhite DA (ed) Drought: a global assessment, vol 1. Routledge, London, New York, pp 3–18 Wobus A, Bleul C, Maassen S, Scheerer C, Schuppler M, Jacobs E, R€oske I (2003) Microbial diversity and functional characterization of sediments from reservoirs of different trophic state. FEMS Microbiol Ecol 46:331–347 Zoppini A (2007) Extracellular enzyme activity in temporary river sediments under drying and rewetting conditions. In: Dick RP (ed) Proceedings of the third international conference, enzymes in the environment: ecology, activity and applications. Viterbo (Italy), 15–19 July 2007, p 29 Zoppini A, Amalfitano S, Fazi S, Puddu A (2010) Dynamics of a benthic microbial community in a riverine environment subject to hydrological fluctuations (Mulargia River, Italy). Hydrobiologia (in press) DOI: 10.1007/s10750-010-0199-6

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Chapter 7

Soil Enzymes as Indication of Soil Quality Ayten Karaca, Sema Camci Cetin, Oguz Can Turgay, and Ridvan Kizilkaya

7.1

Introduction

Soil quality is the phenomenon that has been developed to evaluate the factors effecting soil functionality. It is mainly concerned with sustainable use of soil resources in terms of enhanced agricultural productivity, environmental quality and human health. The reasons for these focusing are rapid growth of the world population, which is demanding an increasing emphasis on sustainable agricultural soil management, and as second ongoing degradation of soil resources, which is also closely associated with the loss of soil quality by climate changes, wildfires, erosion, salinization and agricultural–industrial pollution problems. Early definitions of “soil quality” were actually formalized efforts that had been restricted to analysis of various soil characteristics and evaluation of soil conditions but, in course of time, it has been noticed that analysis of soil properties alone could not be adequate to measure soil quality, as long as the criteria evaluated are associated with the role or function of the soil. Therefore, the term of soil quality has been modified several times related to the ongoing development beyond soil management and the interactions with other ecological systems. For example, in 1980s, soil quality was defined as “the sustained capability of a soil to accept, store and recycle water, nutrients and energy” (Anderson and Gregorich 1984). On the other hand, agricultural activities diffused through much larger ecological systems by time and it has been understood that soil serves many functions not only within or between agricultural lands but also beyond environmental ecosystems.

A. Karaca (*) and O.C. Turgay Faculty of Agriculture, Department of Soil Science, Ankara University, Diskapi, Ankara, Turkey e-mail: [email protected] S.C. Cetin Agricultural Faculty, Soil Science Department, Gaziosmanpasa University, Tokat, Turkey R. Kizilkaya Agricultural Faculty, Soil Science Department, Ondokuz Mayis University, Samsun, Turkey

G. Shukla and A. Varma (eds.), Soil Enzymology, Soil Biology 22, DOI 10.1007/978-3-642-14225-3_7, # Springer-Verlag Berlin Heidelberg 2011

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This development required a revision in the definition of soil quality and a more detailed definition has been stated as “Soil quality is the capacity of a specific kind of soil to function, within natural or managed ecosystem boundaries, to sustain plant and animal productivity, maintain or enhance water and air quality, and support human health and habitation” by Soil Science Society of America (1995). Today, the common framework to evaluate soil quality is generally based on a group of facts mainly including soil functions, processes, attributes, indicators and methodology. In this stepwise evaluation, soil function is the answer of the question of “what the soil does” and soil processes can be defined as the characteristic features of a soil use at anytime. Soil quality attributes describe the measurable soil characteristics that affect the ability of the soil to achieve various functions. In many cases our ability to assess soil quality is complicated because it is not usually possible to directly measure the rate of specific soil processes due to their simultaneity, diverse and conflicting natures and interactions in time, space, and intensity. Therefore soil quality indicators that are associated with specific soil processes can be used as an indirect and useful measure of soil quality changes. In the literature, there have been many attempts to discover possible links between soil quality indicators and specific soil functionality (i.e., plant production) (Doran and Parkin 1994; Gregorich et al. 1994; Larson and Pierce 1994). These works revealed that soil quality indicators could represent either a single or a variety of soil attributes, be easily measured, have some sensitivity to soil management in a wide range, and have also a relatively low sampling error. Supplying available methodology with the convenience of duplication and high accuracy and speed would be also effective factors on the selection of soil quality indicators. Basic soil quality indicators is expected to integrate the combined effects of the soil’s physical, chemical and biological properties and processes, be applicable in many various management and climate conditions, accessible to wide group of users with different expertise and be included by existing soil databases. Soil physical quality is simply an organization of mineral particles, pores and water. The indications of the relationships between these components are usually stated as topsoil depth, bulk density, porosity, aggregate stability, texture, crusting, and compaction and primarily reflect the conditions of root growth, seedling emergence, infiltration, or movement of water within the soil profile (Dexter 2004). The poor soil physical quality implies poor water infiltration, run-off of water from the surface, hard-setting, poor aeration, and poor rootability, while opposite or absence of these conditions is attributed to good soil physical quality. Since the changes in soil physical conditions occur simultaneously, several authors have mentioned that no single parameter of soil physical conditions has been considered as a measure of soil physical quality (Dexter and Czyz 2000), and a range of soil physical properties should be integrated to obtain an overall assessment of soil physical quality (Canarache 1990). The soil quality in terms of soil chemical aspects is related to three main functions in soil, (a) recycling of soil organic matter, (b) storage and gradual release of nutrients, and (c) buffering of potentially toxic elements (Warkentin 1995). These functions actually regulate nutrient flow and the transport of toxic elements

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through soil solution where they are taken up by the plants. Recycling of organic materials depends on the content of mineralisable organic matter and microbial biomass responsible for the continuous supply of nutrients to the soil solution. Function (b) above is controlled by the maintenance of optimum nutrient concentration in the soil solution whereas function (c) needs lower concentrations of toxic elements in the soil solution. The attributes and indicators of the soil’s physical and chemical processes summarized above could be often considered to be adequate to evaluate maninduced changes in soil quality, but on the other hand, they usually represent slowly changing features of soil (i.e., soil structure, organic matter pool and nutrient balance) and since many of the changes in soil physical and physico-chemical conditions take place over the long term, a soil quality evaluation based on these features would require combined analysis of a large group of soil physical and chemical parameters (Gil-Sotres et al. 2005). On the other hand, over the last two decades, there has been an increasing scientific focus on soil biological-biochemical aspects, which have been reported to be more sensitive to the slight modifications pertaining to soil management practices and land use changes (Klein et al. 1985; Powlson et al. 1987; Nannipieri et al. 1990; Ellert and Gregorich 1995; Yakovchenko et al. 1996) and thus may provide more dynamic indications of soil quality changes. Among the indigenous soil biological components, soil microorganisms have a key role in a number of important biochemical processes such as soil microbial activities, responsible for the cycles of bio-elements (C, N, P, and S) and energy transfer in soil ecosystem. Basic source of soil microbial activities is soil organic matter and depending on the land use and other soil characteristics, microorganisms are in a continuous labor to govern soil organic matter and, in most cases like stress conditions caused by adverse anthropogenic effects, this can be rapidly reflected either to the microbial diversity level or to biologically active soil organic matter components i.e., microbial biomass, enzymes and other ephemeral organic compounds i.e., proteins and carbohydrates. Recent developments in the studies of soil microbial diversity have provided an innovative perspective to soil scientists and enabled them to have an insight into the structure and function of soil microbial communities which used to be unknown and hence called as “black box” until 2 decades ago. In relation to the concept of soil quality, the soil microbial community changes can be stressed in two distinct ways (1) verification of the composition and distribution of different functional groups of soil microorganisms and (2) analysis of the dynamics of specific organisms or communities under changing soil conditions (Visser and Parkinson 1992). In other words, a change in entire or specific soil microbial communities could be reckoned as a sensitive indication of a change in soil quality conditions. However, these sensitive tools are usually based on the use of laboring molecular techniques (analysis of intra- and extra-cellular markers i.e., DNA, RNA and cell-wall fatty acids) and generally requires microbiological expertise, costly reagents and wellequipped laboratory facilities. This actually may help to explain increasing attention on soil quality studies focusing on microbial and biochemical soil quality

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indicators and taking general (microbial activity) and specific (hydrolytic enzymes) biochemical parameters into consideration. As a useful tool of soil biochemical quality, soil enzyme activities have been often suggested as sensitive indicators of soil ecological quality because (1) they measure principal microbial reactions involving nutrient cycles in soil, (2) they may easily respond to changes in soil by natural or anthropogenic factors, and (3) they can be easily measured (Gianfreda and Bollag 1996; Calderon et al. 2000; Drijber et al. 2000; Nannipieri et al. 2002). Nevertheless, up until now, the validation of soil biochemical properties including soil enzymes as a universal soil quality criterion has still been under discussion due to the contradictory results obtained from the studies using these properties as an individual parameter or an index component. The reasons for questionable findings on biochemical approaches in the literature have been ascribed to the factors such as different methodological perspectives, the lack of standard analysis and reference values, inadequate databases for high quality soils, high degree of variability between biochemical properties and seasonal or edaphic influences (Gil-Sotres et al. 2005). These drawbacks can certainly be overcome by increasing scientific efforts to understand the behaviors of different soil characteristics and how they function and relate to each other in both disturbed and undisturbed soils. With these in the mind, we will concentrate on the assessment of soil biochemical quality in specific regard to soil enzyme activities. The purposes of this chapter is to emphasis soil enzymes as a soil component, their ecological distribution and behaviours under different environments and finally their potential as a soil quality indicator reflecting ecosystem disturbance.

7.2

7.2.1

Soil Enzymes: Classification, Sources, States, Affecting Factors and Activation or Inhibition Enzyme Classification

Enzymes classification in biological systems is based on reaction type and classified as follows: (1) oxidoreductases: oxidation-reduction, (2) transferases: transfer of functional groups, (3) hydrolases: hydrolysis, (4) lyases: elimination groups for forming double-bond, and (5) izomerases: izomerisation and (6) ligases: forming bond with ATP hydrolysis (Voet and Voet 1995). Soil enzymes mainly belong to hydrolases and the rest of them belong to other classes such as oxidoreductases, transferases, and lyases (Dick and Tabatabai 1992). Location of enzymes is considered another classification of soil enzymes: endocellular or intracellular (in organism) and extracellular (out of organism) (Kunito et al. 2001). Arylsulphatase is an extracellular enzyme whereas dehydrogenase is an intracellular enzyme. Commonly studied soil enzymes can be summarized as: (1) phosphatases (EC 3.1.3.1): it hydroles compounds of oganic phosphorus and

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transforms them into different forms of inorganic phosphorus, (2) urease (EC 3.5.1.5): acting hydrolysis of urea to carbon dioxide and ammonia (3) dehydrogenase: oxidation of organic matter (intracellular), (4) b-glucosidase (EC 3.2.1.21): it catalyzes the hydrolysis of b-D-glucopyranoside and is one of the three or more enzymes involved in the saccharification of cellulose, (5) arylsulphatase (EC 3.1.6.1): involving in the hydrolsis of arylsulphate esters by fission of the oxygen–sulphur (O–S) bond, (6) catalase: involving in microbial oxidoreductase metabolism (intracellular) and may be related to the metabolic activity of aerobic organisms. (7) dehydrogenase: these enzymes are found in all living organisms and take part in many reactions involved in energy transfer in microbial metabolic reactions.

7.2.2

Sources of Soil Enzymes

Sources of soil enzymes are showed in Fig.7.1. Enzymes are originated from mostly microorganisms and as well as plants and animals (Bandick and Dick 1999).

7.2.3

States

State of enzymes has been used to define the phenomenon that enzymes exist in the soil (Dick and Tabatabai 1992). When an enzyme enters into a soil system, enzyme SOIL ENZYMES ABIOTIC ENZYMES 1. Accumulated Enzymes

a. Bound to microbial cellular components b. not associated with cellular components ii. originating from plant roots 2. Continuously released Extracelular enzymes

a. microorganisms

ENDOCELLULAR ENZYMES 1. proliferating microorganisms 2. plant roots 3. soil fauna i. in non-proliferating cells ii. in intact dead cells iii.in cellular fragments i. originating from microorganisms and soil fauna *endocelllular

enzsymes from disrupted cells enzymes

**extracellular

b. plants

Fig. 7.1 Sources of enzymes in soils (Adapted from Dick and Tabatabai (1992) according to Skujins (1978))

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can be found in different states in the soil. These states are: adsorption, microencapsulation, cross-linking, copolymer formation, entrapment, ion-exchange, adsorption and cross-linking and covalent attachment. These mechanisms are considered as the protective effect of soil on extracellular enzyme activity. Adsorption of enzyme by clay minerals is a protective stabilization from microbial attack (Esminger and Gieseking 1942). When free enzymes enter into a soil system, either it can complex with humic colloids or can be stabilized on clay surfaces and organic matter (Boyd and Mortland 1990). Soil enzymes are stabilized by soil organic matter rather than by inorganic components (Dick and Tabatabai 1992). Organic matter protects urease against microbial attack and other processes (Conrad 1940) and enzyme activities are significantly correlated with the organic matter content of soils. In addition, free enzymes can complex with humic acid and these complexes are much more stable to heat and enzymatic degradation than the free enzymes (Serban and Nissenbaum 1986).

7.2.4

Affecting Factors

Enzymatic reactions mostly depend on pH, ionic state, temperature and the presence or absence of inhibitor and activator (Burns 1978; Tabatabai 1982, 1994). In general, every enzyme has an optimum pH value and lower the activity up and below this optimum. Temperature influences enzyme activity in different ways than chemical reactions. Actually, with increasing of temperature every 10
C, chemical reaction rate is doubled but enzymatic reaction rate is increased until optimum pH and then decreased (Tabatabai 1994). Enzymatic reactions are temperature-dependent. However, enzymes are large protein molecules that excess temperature (>50
C) causes of denaturating of protein structure of enzyme (Campbell and Smith 1993). Special cations are required for enzyme activities such as Ni for urease for acting role in active site (Mathews and van Holde 1995). Enzyme activities decrease in the presence of the inhibitor and increase in the presence of an activator.

7.2.5

Enzyme Inhibition and Activation

Enzyme inhibitor can be defined as reducing agent of enzyme activity whereas enzyme activators are stimulating agent of enzyme activity (Voet and Voet 1995). The effects of inhibitor and activator on enzyme are shown in Fig. 7.2. The effects of both agents are mostly on Km. parameter. Km. is defined as “the substrate concentration at which the reaction rate is half maximal” (Stryer 1995). As seen in the figure, Km. values increased in the presence of inhibitor and decreased in the presence of the activator and V.

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Fig. 7.2 The effects of inhibitor and activator on enzyme activity (Voet and Voet 1995)

Enzyme

E+Activator

Vo E+Inhibitor

KmA

7.3

Km

KmI

Indication of Soil Enzymes

Soil enzymes are important in ecosystem functioning with their catalytic feature on nutrient cycling (Bandick and Dick 1999). Soil enzymes have a crucial role in C (b-glucosidase and b-galactosidase), N (urease), P (phosphatases), and S (sulphatase) cycle. Dick and Tabatabai (1992) explain the importance of soil enzymes as “soil enzymes are useful in describing and making predictions about an ecosystem’s function, quality and the interactions among subsystems”. Bandick and Dick (1999) reported that soil enzymes are used as soil quality indicators due to “their relationship to soil biology, ease of measurement and rapid response to changes in soil management”. Using soil enzymes as soil quality indicators can be classified into three distinct areas: (1) pollution indicators, (2) ecosystems perturbations indicators and (3) agricultural practice indicators. Soil is a vital resource not only an environmental filter providing the quality of both water and atmosphere but also being a very complicated medium pursuing sustainability of ecosystems (Trasar-Cepeda et al. 2000). Depending on environmental degradation, soil quality also changes; however, soil quality standards’ has been established later than air and water quality standards (Dick 1997). Soil quality is measured by all soil physical, chemical and biological properties. Microorganisms and biochemical activities have a crucial role in soil ecosystems. Soil enzymes are significant in soil functioning because of the following features: (1) having a role in decomposition of organic inputs, (2) transformation of soil organic matter, (3) releasing nutrients from the unavailable to the available form to plants, (4) participating in N2 fixation, (5) detoxification of xenobiotics (unnatural compounds such as pesticides, industrial wastes, etc.), and (6) participating in nitrification and denitrification processes (Dick 1997). The effects of different type of pollutants on soil enzyme activities are summarized by Dick (1997). Table 7.1 exhibits the sum of his review paper. Generally, heavy metals inhibit soil enzyme activities; pesticides behave differently depending on the type of pesticide either by inhibition/stimulation or have little effect or even no effect. Industrial amendments or contaminants (fly ash, pulp and paper mill effluent) influence positively or negatively soil enzyme activities depending on the

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Table 7.1 Pollution effects on soil enzyme aactivities in literature (summarized from review of Dick (1997), references therein) Pollution type

Studied enzyme

Influence

Heavy metals (Hg, Ag, SP, LG, CL, LA, Cr, Cd) BG

Inhibition

Heavy metals

ARS AC, UR, IN

High inhibition Low inhibition

Pesticides

Enzyme activities

Little or no effect

Insecticides, fungicides, herbicides Pesticides

DH

Stimulation

PH, UR

Slight increase or little effect

Glycophosate, paraquat, Carboryl Atrazine

IN UR, PH UR, IN PH UR, PH, IN

Paraquat, glyphosate Purified Complex with montmorillonite Pesticides (accidental spill) Alachlor (10,000 mg kg1) alone or Mixture atrazine and metochlor Herbicide (imaxethapyr) Pesticides atrazine Herbicide 2,4-D salt formulation 2,4-D isoctyl ester formulation Industrial amendments or contaminants Fly ash Fly ash

UR

Stimulation No effect Inhibition No effect No effect (except very high concentration) No effect Stimulation

Pulp and paper mill effluent Hydrocarbons Jet fuel (50 and 135 mg g1)

Recommended indicator

ARS

References Frankenberger and Tabatabai (1981, 1991a, b), Eivazi and Tabatabai (1990), Deng and Tabatabai (1995) Al-Khafaji and Tabatabai (1979), Bardgett et al. (1994), Yeates et al. (1994) Burns (1978), Ladd (1985) Tu (1981)

Davies and Greaves (1981), Baruah and Mishra (1986), Tu (1992, 1993) Gianfreda et al. (1993).

Gianfreda et al. (1994)

DH ES

Inhibition Inhibition (with herbicide mixture)

Dzantor and Felsot (1991)

DH PR, FDA PH, IN, BG, UR DH, UR DH, UR

Inhibition Stimulation Inhibition Little effect Inhibition

Perrucci and Scarponi (1994) Voets et al. (1974) Rai (1992)

UR, ARS DH

Stimulation Inhibition

McCarty et al. (1994)

PH, SP, DH, IN CT IN, DH

Inhibition Stimulation Stimulation

Pichtel and Hayes (1990)

FDA

Inhibition

IN

Kannan and Oblisami (1990) Song and Bartha (1990)

(continued)

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Table 7.1 (continued) Pollution type

Studied enzyme

Influence

Recommended indicator

References

Atmospheric pollution Acid precipitation (lab. conditions) Acid precipitation (field conditions) Industrial air pollutants (N and S gases) Traffic pollution (heavy metals)

Soil enzymes DH, PH, UR

Inhibition No effect

Bitton and Boylan (1985) Bitton et al. (1985)

CL

Inhibition

Ohtonen et al. (1994)

CL, AML

Inhibition

Joshi et al. (1993)

SP sulphatase, LG L-glutaminase, CL cellulase, LA L-aspariginase, BG b-glucosidase, ARS arylsulfatase, AC acid phosphatase, UR urease, IN invertase, DH dehydrogenase, PH phosphatase, ES esterase, PR protease, FDA 3,6-diacetylfluorescein hydrolysis, CT catalase, AML amylase

type of enzyme. Hydrocarbons, acid precipitation, industrial air pollution (N and S gases) and traffic pollution (heavy metals) adversely affect soil enzyme activities. Soil enzymes are used as pollution indicators by many researchers due to agricultural practices and organic pollution (Gianfreda et al. 2005), irrigation by polluted water (Zhang et al. 2008; Filip et al. 1999; Barton et al. 2000), sewage sludge and municipal waste application (Fernandes et al. 2005; Kizilkaya and Heps¸en 2004; Kizilkaya and Bayrakli 2005; Bastida et al. 2008; Antolin et al. 2005), pesticides application (Gundi et al. 2005; Cai et al. 2007; Fragoeiro and Magan 2008; Yu et al. 2006; Kucharski and Wyszkawska 2008), heavy metals (Kizilkaya et al. 2004; Bhattacharya et al. 2008; Hinojosa et al. 2008; Kunito et al. 2001). Indicators are very important for resource managers in order to understand ecological change (Dale et al. 2008). Dale and Beyeler (2001) summarized the criteria for ecological indicators: (1) easy to measure, (2) sensitive on system stresses, (3) respond to stress, (4) anticipation of change in the ecological system, (5) predict changes, (6) being integrative, (7) ability to respond to natural disturbances, anthropogenic stresses and changes over time, (8) variable with response, (9) having attention of measured parameters spatial and temporal change (Dale et al. 2008). Soil enzymes are used as the ecosystem’s perturbations indicators by many scientists related to changing land use (Sicardi et al. 2004; Acosta-Martinez et al. 2003, 2007; Trasar-Cepeda et al. 2008a; Gil-Sotres et al. 2005; Lin et al. 2004; devegetation and revegetation (Bastida et al. 2006; Izquierdo et al. 2005), forest fires (Staddon et al. 1998; Fioretto et al. 2005; Zhang et al. 2005; Boerner et al. 2008; Camci Cetin et al. 2009), and changing climatic conditions (Sardans and Penuelas 2005; Sowerby et al. 2005). Many authors suggested soil enzyme activities were sensitive, reliable and early indicators for discriminating land use, reflecting soil restoration and fire stress (Sicardi et al. 2004; Izquierdo et al. 2005; Caravaca et al. 2003; Fioretto et al. 2005; Camci Cetin et al. 2009). Farming practices may cause modification in soil environment and providing good conditions for plant growth such as irrigation, tillage, applying fertilizers and

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pesticides, addition of amendments, applying of plant residues, and using crop rotations. All agricultural practices can affect nutrient turnover and microbial activity by influencing soil biological properties (Curci et al. 1997). Soil enzymes play a crucial role in catalytic reactions involved in organic matter decomposition and nutrient cycling (Ajwa et al. 1999). Many studies showed that agricultural practices (crop rotation, mulching, tillage, application of fertilizers, and pesticides) might have different effects on both soil enzymes and microbial activities (Ladd 1985; Dick et al. 1987; Tabatabai 1994). Enzyme activity was found to be the most strongly depressed soil property under intensive agronomic use compared with other biochemical parameters (Saviozzi et al. 2001). Soil enzymes are used as agricultural practices indicators by many researchers due to irrigation (Zhang and Wang 2006), application of fertilizers (Kandeler et al. 1999; Yang et al. 2008; Melero et al. 2007; Bell et al. 2006; Saha et al. 2008a; Marinari et al. 2000; Hu and Cao 2007; Chang et al. 2007; Acosta-Martinez et al. 1999; Patra et al. 2006), application or amendments (Saha et al. 2008b; Martens et al. 1992; Rajashekhararao and Siddaramappa 2008; Leon et al. 2006), different management and farming systems (FlieBbach et al. 2007; Melero et al. 2008; Gajda and Martyniuk 2005; Monokrousos et al. 2006; Sarapatka 2002; Benitez et al. 2006; Landgraf and Klose 2002; Gajda and Martyniuk 2005; Kremer and Li 2003; Bandick and Dick 1999), crop rotation (Benintende et al. 2008; Dodor and Tabatabai 2005, 2003), and tillage (Mina et al. 2008; Ekenler and Tabatabai 2004, 2003; Acosta-Martinez and Tabatabai 2001; Madejon et al. 2007; Curci et al. 1997; Deng and Tabatabai 1997). Trasar-Cepeda et al. (2000) indicated that soil enzymes as indicators of soil pollution were limited due to their ability for reflecting soil degradation caused by pollution. Doran and Parkin (1994) and Elliott (1997) suggested that pollution indicators should have the following features: (1) pollutant sensitivity, (2) reflecting ability of different levels of pollution, (3) response either increases or decreases, (4) sensitivity of different pollutants, (5) discriminating between pollutant effect and prior degradation of the polluted soil and (6) differentiating all pollutants based on different degrees of soil degradation they cause (Trasar-Cepeda et al. 2000 and references therein). For that reason, many authors (Beck 1984; Stefanic 1994; Perucci 1992; Sinsabaugh 1994; Stefanic 1994; and Yakovchenko et al. 1996) suggested that using soil enzyme activities as indicators of soil contamination would be acceptable using soil enzymes and other biochemical properties in combination to develop more complex expressions for minimizing limitation of soil enzyme as a pollution indicator (Trasar-Cepeda et al. 2000 and references therein).

7.3.1

Soil Enzymes as Pollution Indicators

Intensive agricultural practices and organic pollution also change soil enzyme activities. Gianfreda et al. (2005) found that arylsulfatase, b-glucosidase, phosphatase, urease, dehydrogenase, and fluorescein diacetate hydrolase were lower or had

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no activity in non-cultivated soils (heavily or moderately polluted by organic contaminants) when compared to agricultural soils. Irrigation systems influence soil systems depending on irrigation water quality. Long-term sewage irrigation resulted in an increase of heavy metal quantity in soil and caused a positive impact on the microbial community and soil quality (Zhang et al. 2008). Similarly, soil enzyme activities (b-glucosidase, b-acetyl-glucosaminidase, proteinase, and phosphatase) and microbial biomass were greater in wastewater irrigated soils than in wastewater unirrigated soils (Filip et al. 1999). Denitrifiers and denitrification enzyme activity were higher in wastewater-irrigated soils than wastewater-unirrigated soils, although both the types of soils had the same initial denitrifiers and denitrification enzyme activity in forest soil (Barton et al. 2000). Sewage sludge application in agricultural areas is preferred due to containing valuable nutrients and improving soil fertility (Antolin et al. 2005). Addition of sewage sludge increased basal respiration, microbial biomass, metabolic quotient and enzyme activities and their values were positively correlated with increased sewage sludge doses (Fernandes et al. 2005). Kizilkaya and Heps¸en (2004) showed sewage sludge amended soil have higher enzyme activity such as urease, phosphomonoesterase, arylsulphatase than unamended soil. They found that amended sewage sludge might have quickly decomposed resulting in a higher enzyme activity in soil, and sewage sludge treatment had stimulated microbial production of enzyme activity or had more of these enzymes accessible to substrate. Effects of adding different doses (0, 100, 200, and 300 t/ha dry weight) and C/N ratios (3:1, 6:1 and 9:1) of the sewage sludge on activities of b-glucosidase, alkaline phosphatase, arylsulphatase and urease in a clay loam soil at 25
C and 60% water holding capacity were studied by Kizilkaya and Bayrakli (2005). Nitrogen was added in the form of (NH4)2SO4 solution to the sludge to obtain different C/N ratios. Rapid and significant increase in the soil enzymatic activity has been noted at different doses and C/N ratios of the sewage sludge amendments as compared to unamended ones. Enzyme activities varied with differences in incubation period. Soils with the highest C/N ratio and sludge dose had the highest b-glucosidase activity. Alkaline phosphatase and aryl sulphatase showed an increment in their activity during the first 30 days of incubation followed by a pronounced decrease compared to unamended soil. Urease activity, however, showed an increase within 15 days, and thereafter activity declined. The highest activities of urease, alkaline phosphatase and arylsulphatase were observed in soil amended with a low C/N ratio and the highest dose of sludge. Another type of waste usage in agricultural soils is solid municipal waste application. Soil enzyme activities (b-glucosidase, urease, alkaline phosphatase, and o-diphenol oxidase; humus-associated enzymes) increased with the dose of application of solid municipal waste at different levels (low, medium, high, and very high) but only a certain level, after this level either sustain the same level or decreased (Bastida et al. 2008). Both natural events and anthropogenic activities continuously influence soil quality (Puglisi et al. 2006). Application of sewage sludge promotes soil biological activity (Saviozzi et al. 1999). Basal respiration, microbial biomass and enzyme activities (urease, BAA-protease, phosphatase, and b-glucosidase) under barley

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cultivation, were improved by repeated sewage sludge application (Antolin et al. 2005). Pesticides application in agricultural system affects soil enzyme activities in different ways depending on type, concentration, application duration, number of pesticides and other amendments in soils. Also, different soil enzymes response differently either increase or decrease. Insecticides (monocrotophos, quinalphos, and cypermethrin in single or combination) significantly increased dehyrdogenase activity but decreased in the highest concentration (Gundi et al. 2005). Dehydrogenase activity was decreased because of high residual acetochlor (Cai et al. 2007). However, application of organic fertilizer in soils treated by acetochlor resulted in stimulation of dehydrogenase activity in soil. Addition of pesticides (simazine trifuralin, and dieldrin; inoculation of soil microcosms with Trametes versicolor and Phanerochaeto chrysosporium) in soil decreased dehyrdogenase activity in most treatments (Fragoeiro and Magan 2008). Acid phosphatase, alkaline phosphotase, urease, catalase, and invertase activities inhibited only first and second pesticide application (chlorotholanil) and especially urease, catalase and acid phosphatase were affected (Yu et al. 2006). Kucharski and Wyszkawska (2008) found that urease and dehydrogenase were the least tolerant and alkaline phosphatase was the most tolerant to the effect of herbicide (Apyros 75 WG). The effect of heavy metals was evaluated many authors on soil enzyme activities. Studies concerning the effects of heavy metals in soils were shown in Table 7.2. Heavy metals are known to cause long-term toxic effects within ecosystems and can have a negative influence on soil enzymatic processes (Kizilkaya et al. 2004). They can also affect microbial proliferation and enzyme activities by masking catalytically active groups, altering protein conformation or competing with other metals involved in the formation of enzyme–substrate complexes (Eivazi and Tabatabai 1990). Fluorescein diacetate, b-glucosidase, urease, phosphatase, and arylsulphatase were negatively correlated with soils metals (Cd, Cr, Cu, and Pb) in

Table 7.2 Studies concerning the effects of heavy metals in soils (summarized from the study of Puglisi et al. (2006), references therein) Treatment or application Enzyme References Copper contamination mine and arable land BG, PH, UR Leiros et al. (1999) Metal contaminated mine ARS, PH, UR, DH, Majer et al. (2002) XY, PR Bioremediation monitoring BG, PH, UR, DH, PR Pascual et al. (2000) Metal-contaminated grassland BG, PH Kuperman and Carriero (1997) Cadmium and sludge PH, DH, AOA Dar (1996) Tanning and landfill effluent, hydrocarbon ARS, BG, PH, UR, Trasar-Cepeda et al. contaminated DH (2000) Bioremediation monitoring UR, DH, CT, LP Margesin et al. (2000) Heavy metal contamination ARS, BG, PH, UR, Hinojosa et al. (2004) DH BG b-glucosidase, ARS arylsulfatase, UR urease, DH dehydrogenase, PH phosphatase, PR protease, CT catalase, XY xylanase, AOA arginine ammonification activity, LP lipase

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long-term irrigation of sewage treatments (Bhattacharya et al. 2008). The highest enzyme activities were found in non-polluted soils compared with polluted but restored and polluted but un-restored (Hinojosa et al. 2004). Acid phosphatase, alkaline phosphatase, arylsulfatase, cellulose, dehydrogenase, protease, urease, and invertase (except b-D-glucosidase) increased with addition of sewage sludge or compost; however, the ratio of enzyme activities to microbial biomass showed a decreasing trend with these applications (Kunito et al. 2001).

7.3.2

Soil Enzymes as Ecosystems Perturbations Indicators

Changing land use from one type to another type generally affects the soil ecosystems. Land use conversion from natural grazed pastures to commercial Eucalyptus grandis plantations, showed that no significant effect was found on the number of cellulolytic aerobes, P-solubilizers and Azotobacter ssp. communities, whereas significant effect on soil respiration, the C-mineralization coefficient, dehydrogenase, fluorescein diacetate hydrolysis and acid and alkaline phosphatase activities (Scardi et al. 2004). Scardi et al. (2004) reported that land use and management practices alter the total amount and composition of soil organic matter (Reaves 1997) and significantly change the enzyme activities (Dick 1997). Natural systems and distributed systems are different from their biological activities. b-glucosidase and acid phosphatase, and arylsulfatase (except a-glucosidase and b-glucosaminidase) based on land use showed the following orders, respectively: pasture > forest > agriculture and forest ¼ pasture > agriculture (Acosta-Martinez et al. 2007). Land use type and the selected enzyme kind respond in different ways. Land use changed the level of organic matter and enzyme activities (acid phosphomonoesterase, b-glucosidase, phosphodiesterase, arylsulfatase, urease, protease, hydrolyzing bezoyl-argininamidase, invertase and carboxymethyl-cellulase) based on the type of land use (agricultural and forest) and type of enzyme (Trasar-Cepeda et al. 2008a). Although biomass-C, urease and b-glucosidase were used as possible indicators of land use on soil (Gil-Sotres et al. 2005), land use did not always modify biochemical properties in the same way and no biochemical property can be used as an universal indicator for discriminating land use (Trasar-Cepeda et al. 2008a). Dehydrogenase activity was lower in plastic-greenhouse vegetable cultivation compared with rice-wheat rotation; however, lower activities (urease, invertase, and phosphatase) were found in open-field vegetable cultivation soils than rice-wheat rotation soils (Lin et al. 2004). Excessive use of N and P fertilizers can be harmful to long-term soil quality due to acidification, nutrient enrichment and influencing microbial activity especially influenced by land use patterns. Natural systems changed to agricultural systems and not only vegetation but also soil biological properties altered in soil ecosystems. The higher enzyme activities (b-glucosidase, b-glucosaminidase, arylamidase, alkaline, and acid phosphatase, phosphodiesterase, and arylsulfatase) were found in the conservation reserve

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program, native grassland and rotation with other crops (wheat or sorghum) compared with continuous cotton. The results showed that crop rotation provided higher enzyme activities in soils (Acosta-Martinez et al. 2003). Devegatation and revegetation influence soil quality compared with undistributed soils. Dehydrogenase and protease activities were lower in devegetated soil (devegetation of Pinus halepensis and natural shrubs) than undistributed soil (Bastida et al. 2006). Elimination of vegetation caused a long-term negative influence on the biochemical state and microbial activity of soil. Soil quality has not been recovered even after 15 years due to deforestation. Protease and b-glucosidase activities were higher in revegetation with Casuarina equisetifolia than with Anacardium occidentale; however, urease, protease, acid phosphatase, and b-glucosidase activities were significantly greater in revegetated soils (soil restoration: after revegetation of a mining area) than in the bare soil 4 years after planting (Izquierdo et al. 2005). Forest fires are considered as natural disturbances in forest ecosystem (GonzalezPerez et al. 2004; Zhang et al. 2005) and cause the most dramatic changes in forest ecosystems (Shakesby et al. 2007). Due to the low volatilization temperature of N, most of the nitrogen found in biomass and soil is lost to the atmosphere when forest fires occur. After a forest fire, many complex influences occur in the forest ecosystems. These influences can be summarized into two main groups: (1) reduction of biomass and (2) alteration of below-ground quality–quantity and functionality (Zhang et al. 2005; Neary et al. 1999). Many studies have showed the effect of forest fire on soil enzyme activities (Zhang et al. 2005; Ajwa et al. 1999; Boerner and Brinkman 2003; Boerner et al. 2000, 2005; Saa et al. 1993, 1998; Senthilkumar et al. 1997). Hart et al. (2005) reported that soil biological properties are more sensitive than the other soil properties to soil heating because of their low fatal temperature (<100
C) (DeBano et al. 1998). Only some enzyme activities discriminate the fire effect on forest ecosystems as bio-indicators. Among soil enzymes (cellulose, xylanase, invertase, trehalase, and protease), only invertase activity decreased in burned plots throughout the study period (Fioretto et al. 2005) There was no differences between unburned (Quercusdominated forests) and burned sites based on soil enzyme activities(acid phosphatase, a-glucosidase, phenol oxidase, chitinase and L-glutaminase) sampled in early spring and late spring/early summer; however, there was a clear separation of burned and unburned soils due to L-glutaminase activity in late summer/early autumn (Boerner et al. 2008). Acid phosphatase activity could be used for evaluating fire effects on soils compared with alkaline phosphatase and arylsulfatase activities (Staddon et al. 1998). Different soil enzymes were investigated by different researchers for discriminating fire stress on soil quality. Some activities were decreased and others were increased. Invertase and proteinase activities were declined by burning but acid phosphatase, polyphenoloxidase, and peroxidase activities were increased (Zhang et al. 2005). Phenol oxidase activity was decreased by the fire effect relative to control (Boerner et al. 2008). Urease activity declined by the fire effect and this

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negative effect on urease activity was time-dependent and recovered after 12 years of burning (Camci Cetin et al. 2009). Climatic conditions influence all living things as well as soil quality. The reduction of 10% of soil moisture decreased urease (10–67%), protease (15–66%) and b-glucosidase (10–80%) activities while decreasing of 21% of soil moisture declined urease (42–60%), protease (35–54%), b-glucosidase (35–83%) and acid phosphatase (31–40%) activities and no significant influences were found on alkaline phosphatase activity (Sardans and Penuelas 2005). N-cycling enzymes (protease and urease) were the most influenced by drought. The effect of drought on soil enzyme activities (glucosidase, sulfatase, phosphatase, and leucine amino peptidase) was higher and more pronounced in the Northern European sites than in the Southern European due to having moisture limited feature (Sowerby et al. (2005). The effect of temperature enhanced soil enzyme activities in all sites (Denmark, UK, The Netherlands and Spain); however, Northern Europe soils were more sensitive to changes in rainfall regimes than more moisture limited Southern European soils.

7.3.3

Soil Enzymes as Agricultural Practices Indicators

One of the agricultural practices is irrigation that provides adequate moisture level in soil for plant growth. Zhang and Wang (2006) investigated the impact of subsurface irrigation (10, 16, 25, 40, and 63 KPa) on phosphatase, urease, and catalase activities under tomato cultivation in a greenhouse experiment. Phosphatase and catalase activities increased in more frequent irrigation (10 and 16 KPa) and urease activities decreased under irrigation. Fertilization of soils are conducted in soils by using different fertilizers such as mineral, manure, green manure, compost, and vermicompost. Kandeler et al. (1999) showed that farmyard manure enhanced microbial biomass, urease, deaminase, and alkaline phosphatase activity in soils compared with other treatments (mineral fertilizers) under crop rotations. The stage of plant growth should be the cause of concern on enzyme activities in soils in terms of evaluating the impact of fertilities. Similarly, soil enzyme activities (phosphatase, catalase, invertase, and urease) under different fertilizers (no fertilization, organic manure, organic manure þ N, organic manure þ NP, organic manure þ NK, and organic manure þ NPK) were lower in the early growth stages of cucumber, but enhanced in the late stages (Yang et al. 2008). Also, the type of mineral fertilizers used influences soil enzyme activity depending on the soil enzyme involved in which there is, nutrient cycling (N, P, C, and S). Soil enzyme activities were inhibited by N fertilizer while they were increased by P and K fertilizers. Decrease of urease activity could be explained by activation of nitrification and denitrification causing suppression in urease production (Aon et al. 2001).

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Soil quality demonstrates agroecosystem sustainability and productivity (Melero et al. 2007). Using organic manures rather than chemical fertilizers in crop production is assumed to be a more sustainable type of agriculture (Chang et al. 2007). Excessive use of mineral fertilizer can decline soil organic matter as well as soil fertility (Melero et al. 2007). Thus, application of organic fertilizers becomes favorable for environmentalists, agriculturists and consumers (Chang et al. 2007). Compost fertilized plots (vegetal compost and animal compost) showed increase in enzyme activities (dehydrogenase, protease, glucosidase, and alkaline phosphatase) under dryland agriculture system (Melero et al. 2007). Bell et al. (2006) found increased phosphates activities following manure application likely due to enhanced P mineralization. Dehyrogenase, cellulose, protease, urease, acid phosphatase, and alkaline phosphatase activities (except urease) increased with application of manure; however, acid and alkaline phosphatase activities were adversely affected by chemical fertilizer treatment (Saha et al. 2008a). Dick (1997) reported that phosphatase inhibition was related to the level of PO4 (high level) in soil solution (Chunderova and Zubets 1969) and phosphate may inhibit phosphatase synthesis and orthophosphate was known as a competitive inhibitor of acid and alkaline phosphatase activity (Juma and Tabatabai 1978). Animal manure enhances soil enzyme activities than mineral fertilizers and soil microbial and biochemical activities (Deng et al. 2006). Organic fertilizers can be used in agricultural systems especially in organic farming. Compost application is important for establishing and maintaining soil organic matter to a certain level in organic farming (Chang et al. 2007). Organic treatments (sludge vermicompost and stabilized dairy manure) enhanced soil biological activity (microbial biomass and acid phosphatase, dehydrogenase, and protease BAA) due to an enrichment of soil organic matter (Marinari et al. 2000). Hu and Cao (2007) showed that alkaline phosphatase and urease activities were significantly greater in compost systems than in chemical fertilizers and control systems. Chang et al. (2007) found that soil enzyme activities (dehyrogenase cellulose, b-glucosidase, protease, urease, arylsulfatase and acid and alkaline phosphatase) and as well as other microbial properties increased significantly in the compost-treated soils compared with chemical-fertilized soils; however, no significant elevation was observed in studied enzyme activities after the compost dose of 540 kg N ha1 year1. The combination of mulch and fertilizers stimulate enzyme activities compared with fertilizers alone. b-glucosidase and dehydrogenase activities increased with increasing of leaf and N fertilization rate (Acosta-Martinez et al. 1999). Combined used of green manure and nitrification inhibitor increased content of mineral-N (NH4þ and NO3); however, dehydrogenase and nitrate reductase activities decreased with combined application (Patra et al. 2006). Using organic manures and crop residues is recommended in sustainable soil management due to providing maximum benefits and high environmental quality (Rajashekhararao and Siddaramappa 2008). Dehyrogenase activity was higher in composted cattle manure (44–200%) and vermicompost (22–108%) than in control (Saha et al. 2008b) Saha et al. (2008b) concluded that: (1) organic amendments

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applications enhanced organic matter contents and microbial biomass and this provided better potential for higher enzyme production and greater enzyme activities, (2) additions of organic amendments showed different responses on soil enzyme activities depending on organic amendment types, and (3) addition of organic amendments (either cattle manure compost or vermicompost) improved soil quality, increasing soil organic matter content and stimulating biological and biochemical properties. Different kinds of organic amendments originated from animal or vegetal sources can be used in agricultural systems. Organic amendment residues (poultry manure, sewage sludge, barley straw and green alfalfa) applications provided higher enzyme activities (phosphomonoesterases, arylsulfatase, N-acetyl-b-glucosaminidase, b-glucosidase, b-glactosidase, invertase, dehyrogenase, amidase, and urease) in the first year of experiment; however, following subsequent additions did not maintain high enzyme activities (Martens et al. 1992). The most effective amendment increasing soil enzyme activities (except urease) was the straw amendment. Organic amendments influence soil microbial and biochemical properties in different ways depending on the nutrient content. Application of higher rates of organic amendments (rice residues and tree litters: high C content) was favorable soil quality parameters (microbial biomass C, microbial quotient, urease and acid phosphatase activities) (Rajashekhararao and Siddaramappa 2008). Although they evaluated the other soil health parameters (extraction yield of humus and composition of humus) microbial quotient was the most sensitive indicator for reflecting the decline in soil quality. The authors reported that microbial biomass, microbial quotient and soil enzyme activities are used for measuring soil quality as parameters (Doran and Zeis 2000). Addition of different C sources enhanced urease activity at different levels of elevation depending on N levels (Rajashekhararao and Siddaramappa 2008). Higher level of N stimulated urease activity in different C sources. Organic amendments can be used for suppression of plant diseases. Root rot severity was strongly adversely correlated with total C, arylsulfatase and b-glucosidase activities (Leon et al. 2006). b-Glucosidase was not accepted as an useful indicator of disease suppression because it varied over time. Arylsulfatase was the best indicator for reflecting disease suppression. They indicated that applying organic amendments to soils can cause disease suppression by enhancing antagonist microorganisms (Cook 1990) and microbial biomass and activity can be related to microbial competition with pathogens. Conventional agriculture can cause to decrease soil organic matter and reduction in soil fertility (Melero et al. 2008). In terms of this view, sustainable agricultural management practices are crucial in the production of food and human nutrition (Niemi et al. 2008). Conventional cropping systems is required to use mineral fertilizers and pesticides for high crop yield; whereas organic cropping systems is preferred to use plant residues, green manure, farmyard manure and plant rotation for adequate crop yield and high soil quality (Niemi et al. 2008). Inorganic systems, plant production depends primarily on nutrient cycling in soils that are controlled by microbes and their activities and soil enzymes (Monokrousos et al. 2006).

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Different farming systems may change soil parameters especially soil microorganisms and soil enzymes. FlieBbach et al. (2007) investigated the impact of different farming systems [organic: bio-dynamic (farmyard manure compost þ slurry), and bio-organic (farmyard manure þ slurry); conventional: stacked farmyard manure þ slurry, and mineral: mineral fertilizers; and unfertilized] on dehyrogenase activity on soil microbial biomass under 7 year crop rotation. Dehyrogenase activity was lower 39–42% in conventional and 62% in mineral farming systems compared with organic (bio-dynamic) farming system. Transition of farming system (conventional through organic) affects soil enzyme activities. Protease and phosphatase activities increased in organically fertilized soils (Melero et al. 2008). Similarly, arylsulfatase, phosphomonoesterase and esterase activities were greater in organic managed systems than in conventional systems and application of peat amendments increased phosphmonoesterase, phosphodiesterase, leucine aminopeptidase, chitinase, cellobiosidase, a-glucosidase and esterase activities, but decreased arylsulfatase and alanine aminopeptidase (Niemi et al. 2008). Sarapatka (2002) evaluated the effect of different farming system (conventional þ no animal manure: CN, organic þ no animal manure: OA, organic þ no animal manure þ standard cultivation: ONS, and organic þ no animal manure þ minimum soil management: ONM) on acid and alkaline phosphatase activities. The order of acid and alkaline phosphatase activities was: ONM > OA > CN > ONS. The author suggested that soil phosphatase activity was directly related to organic matter content and was affected by farming activities. Monoculture cultivation causes different response on soil enzyme activities as well as conventional crop management compared with organic farming system. Dehyrogenase, acid and alkaline phosphatase activities and microbial biomass C and N levels were lower under monoculture system than under organic and conventional-short rotation systems (Gajda and Martyniuk 2005). Changing farming system to organic farming, especially transition period influences soil biochemical properties. After conversion to farming system from conventional to organic, L-asparaginase, L-glutaminase, urease, acid and alkaline phosphatase were greater under organic management system than conventionally managed asparagus fields (Monokrousos et al. 2006). The experiments were designed as organic farming lasted 2, 3, 5, and 6 years. Acid phosphatase activity increased from the newest to oldest organic managed areas. Many authors evaluated the impact of different management systems on soil quality parameters. Higher enzyme activities (dehydrogenase, o-diphenol oxidase, b-glucosidase, and phosphatase) were found under organic management system compared with conventional and integrated systems (Benitez et al. 2006). The authors suggested that using synthetic chemicals caused the reduction of biochemical activity in soils. Different management systems can affect soil quality the way of soil perturbation. Greater enzyme activities (b-glucosidase and L-asparaginase) were found under succession fallow compared to agriculture and forestry systems, and this elevation could be explained by the lack of tillage that led to higher microbial and

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biochemical activities under successive fallow system (Landgraf and Klose 2002) Similarly, Gajda and Martyniuk (2005) found substantial disturbances in microbial activity of monoculture soil and higher enzyme activities (dehydrogenase and phosphatases) and microbial biomass C and N contents in organic and conventional systems. Declined populations (biomass and respiration rate) and activity of microorganisms in the monoculture soil might cause increasing deleterious microorganisms or decreasing nutrient availability to crops. Fluorescein diacetate hydrolase, dehydrogenase and phosphatase activities were the highest in the native-prairie ecosystem, organic farming, integrating cropping (no-tillage and crop rotation), crop rotation systems than compared with other systems (conventional þ monoculture and conventional þ high agrichemical input) and greater enzyme activities were found in higher weed-suppressive activity, organic matter content and water-stable soil aggregates (Kremer and Li 2003). Bandick and Dick (1999) showed that higher enzyme activities were found in the continuous grass and continuous pasture than in cultivated fields and the highest activity was observed in cover crops plots. Due to monoculture systems decreased soil microbial activities,sustainable agricultural management requires crop rotation. Microbiological and biochemical properties (microbial biomass C and N, metabolic quotient, urease and fluorescein diacetate hydrolysis) were sensitive variables to determine soil rotations’ effects and they could be used as soil quality indicators (Benintende et al. 2008). Also, microbial parameters were more effective management indicators in soil quality than biochemical parameters and all of them should be used as bioindicators when considering the impact of management on soil quality. Trasar-Cepeda et al. (2008b) showed that intensive agricultural system (with crop rotation) has led to low hydrolytic enzyme activities (acid phosphatase, b-glucosidase, phosphodiesterase, arylsulfatase, urease, BAA-protease, invertase, and carboxymethylcellulase) and organic matter content compared to climax soils (oak forest soil). The influence of crop rotation [NERC site: continuous soybean (SSSS), continuous corn (CCCC), corn-soybean (CSCS), corn-corn-oat-alfalfa (CCOM); and CWRC site: CCCC, CSCSC, CCOM, and COMM] and fertilization (0 and 180 kg N ha1) on soil enzyme activities (a- and b-glucosidases and a- and b-galactosidases) were studied by Dodor and Tabatabai (2005). The results showed that : (1) glycosidases activities were significantly influenced by crop rotation but not by nitrogen application at the two sites; (2) greater activities were found in meadow or oat plots and the lowest in continuous corn or soybean plots; and (3) monocropping suppressed glycosidases activities whereas multicropping stimulated them. Sincedifferent cropping systems change soil environment, a soil enzyme should be chosen that is directly involved in soil organic matter cycle because a more accurate way is the measurement of changes than measurement of C and N (Dodor and Tabatabai 2005). Multicropping systems enhanced amidohydrolases activities: however, monocropping systems reduced amidohydrolases activities and the mineralization of organic N in soil was provided by microorganisms through enzyme activities and amidohydrolases activities can be used to anticipate N mineralization and monitor soil quality (Dodor and Tabatabai 2003). Microbiological properties

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are accepted as more sensitive and more responsive indicators to agronomic treatments relative to soil physical and chemical properties as soil quality indicators (Biederbeck et al. 2005). Tillage application may change soil quality through altering soil physico-chemical, hydrological (Rahman et al. 2008), microbiological and biochemical properties and thus influences soil microbial communities and the production of soil enzymes (Acosta-Martinez and Tabatabai 2001). Tillage affects soil nutrient levels and its availability (Etena et al. 1999), distribution of organic matter in the soil profile (Kandeler et al. 1999), soil water and oxygen content (Curci et al. 1997), and soil fertility. Tillage especially influence soil organic matter by exposing more soil organic matter to microbial attack (Mina et al. 2008) and finally rapid loss of soil organic matter (Madejon et al. 2007). Losing soil organic matter causes decline of soil biological activity and crop productivity and increase soil erosion (Madejon et al. 2007). To sum, tillage causes a great perturbation in soil environment. Many researchers conducted the impact of tillage on soil quality parameters as well as soil enzymes. Dehydrogenase activity increased under continuous zerotillage practice and alkaline phosphatase and protease activities were higher in the zero-tillage system over conventional practice; however, cellulose activity was greater in conventional practice compared with other management (Mina et al. 2008) Ekenler and Tabatabai (2004) found that L-glutaminase was the most sensitive N cycling enzyme followed by L-asparaginase, amidase, arylamidase, urease and L-aspartase for discriminating the effect of liming and tillage amendments (notill, ridge-till, and chisel-plow). Also, the most sensitive enzymes reflecting soil management practices were b-glucosidase and b-glucosaminidase whose activities could be accepted early and reliable indicators of changes in soil properties affected by liming and tillage systems (Ekenler and Tabatabai 2003). The combined effect of tillage and residue placements alters soil enzyme activities. Arylamidase activity was highly influenced by tillage and residue placements and the greatest arylamidase activity was found in treatments of chisel/mulch, moldboard plow/mulch, and no-till/double mulch whereas the lowest activity was observed in treatments of moldboard plow/normal and no-till/bare (Acosta-Martinez and Tabatabai 2001). Dehydrogenase, alkaline phosphatase, b-glucosidase, and urease were higher under conservation tillage than under traditional tillage (Madejon et al. 2007). Soil enzyme activities are accepted early and are more reliable indicators than soil physico-chemical properties under different tillage systems. Curci et al. (1997) evaluated the influence of conventional tillage systems (shallowing plowing: 20 cm, deep plowing: 40 cm and scarification: 50 cm) at different depths (0–20, 20–40, 40–50, and 50–70 cm) on soil enzyme activities (acid phosphatase, alkaline phosphatase, phosphodiesterase, pyrophosphatase, arylsulfatase, dehydrogenase, a- and b-glucosidase, a- and b-glalactosidase, urease and nitrate reductase). The results showed that: (1) glycosidase, galactosidase, nitrate reductase, and dehydrogenase activities were influenced by tillage systems, (2) their activities were greater in shallow plowing and scarification than deep plowing plots in the upper layer (0–20 cm) of soils, and (3) no significant differences were found in soil physicochemical properties under different tillage systems.

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No-till systems provide better enzyme activities in soils. Deng and Tabatabai (1997) showed that acid phosphatase, alkaline phosphatase, phosphodiesterase, inorganic pyrophosphatase and arylsulfatase were significantly higher in no-till/ double mulch than in other treatments (no-till/bare, no-till/normal, chisel/normal, chisel/mulch, moldboard/normal, and moldboard/mulch).

7.4

Conclusion

Soil enzymes are important soil components that are closely associated with physicochemical and biological characteristics of soil. However, human activities, agricultural practices and environmental pollution severely influence their existence and activities in soil. Intracellular enzymes are more susceptible to anthropogenic activities. However, extracellular enzymes held in clay and organic components have no linkage with the microorganisms by which it had been previously synthesized and thus their responses are usually slow and permanent while those of intracellular enzymes are more sensitive to the changes in soil environment. On the other hand, it is likely that extracellular enzymes would provide reliable results in the evaluation of long-term impacts of human activities on soil. As a conclusion, depending on their origin, soil enzymes are powerful tools applied in the assessment of short- or long-term changes in soil. Majority of the anthropogenic or environmental factors affecting soil quality can be addressed to the changes in the soil’s physical and chemical aspects, but at the same time they provoke the changes in soil enzyme pool as an indication of soil biological quality, meaning that overall soil quality parameters are closely related to each other. For example, agricultural practices such as organic matter applications, irrigation, fertilization and tillage change soil aggregation processes, nutrient recycling and also soil biological activities governing soil enzyme production. The aggregation of soil mineral particles is one of the major soil processes that is known to develop subsequently the microbiological–biochemical activities. Both aggregation and disaggregation processes are actually long-term events, but on the contrary, changes in soil microbiological–biochemical conditions are fast and occurs in a short span of time. Soil enzymes that substantially originated from soil microorganisms may therefore be more preferable compared to soil physical and chemical quality parameters. For example, the index values of structure stability, erosion rate and erodibility are common soil criterion applied for the determination of soil erosion sensitivity, but in most cases these parameters are time-consuming and economically not reasonable. On the other hand, extracellular soil enzymes can be considered as an indication of soil erodibility (Kizilkaya et al. 2003 and they also enable researchers efficiently, to monitor the impacts of a wide group of agricultural management practices and the differences between remediated and severely degraded soil conditions. Due to these advantages over soil physical and chemical quality attributes,

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soil enzyme activity is considered to be an integral index of soil health but it should be kept in mind that an accurate evaluation of soil health needs to be complemented by physical, chemical and other biological parameters. Although many scientific communities have recognized the potential value of soil enzyme activities as a proper indication of soil quality, a number of problems i.e., Differences in methodological perspectives and the lack of standard methods can complicate the interpretation of soil enzyme activities. The variability in soil enzyme values is also related to the fact that natural changes in climatical conditions and soil characteristics occurs simultaneously with the changes resulted from agricultural activities or environmental pollution. This actually makes more difficult to answer the question of which factor would virtually operate soil biochemical processes and to compare soil enzyme data obtained from different regions or climate zones. Nevertheless, soil enzyme activity is a fundamental indication of most changes in soil condition and hence soil quality, but it seems necessary to develop site-specific evaluation criteria for more accurate use of soil enzyme activities as a soil quality parameter.

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Sicardi M, Garcia-Prechac F, Frioni L (2004) Soil microbial indicators sensitive to land use conversion from pastures to commercial Eucalyptus grandis (Hill ex Maiden) plantations in Uruguay. Appl Soil Ecol 27:125–133 Sinsabaugh RL (1994) Enzymic analysis of microbial pattern and processes. Biol Fertil Soils 17:69–74 Skujins JJ (1978) History of abiotic soil enzyme research. In Soil Enzymes. R.G. Burns (ed). Academic Pres. New York, pp 1–49 Soil Science Society of America (1995) Statement on soil quality. Agronomy News, June. Song HG, Bartha R (1990) Effects of jet fuel spills on the microbial community of soil. Applied and Environmental Microbiology 56:646–651 Sowerby A, Emmett B, Beier C, Tietema A, Penuelas J, Estiarte M, Meeteren MJMV, Hughes S, Freeman C (2005) Microbial community changes in healthland soil communities along a geographical gradient: interaction with climate change manipulations. Soil Biol Biochem 37:1805–1813 Staddon WJ, Duchesne LC, Trevors JT (1998) Acid phosphatase, alkaline phosphatase and arylsulfatase activities in soils from a jack pine (Pinus banksiana Lamb.) ecosystem after clear-cutting prescribed burning and scarification. Biol Fertil Soils 27:1–4 Stefanic G (1994) Biological definition, quantifying method and agricultural interpretation of soil fertility. Romanian Agric Res 2:107–116 Stryer L (1995) Biochemistry, 4th edn. WH Freeman and Company, New York, pp 181–206 Tabatabai MA (1982) Soil enzymes. In: Page AL, Miller RH, Keeney DR (eds) Methods of soil analysis. Part 2. Agronomy 9, 2nd edn. American Society of Agronomy, Madison, WI, pp 903–947 Tabatabai MA (1994) Soil enzymes. In: Page AL, Miller RH, Keeney DR (eds) Methods of soil analysis. American Society of Agronomy, Madison, WI, pp 775–833 Trasar-Cepeda C, Leiros MC, Seoane S, Gil-Sotres F (2000) Limitations of soil enzymes as indicators of soil pollutions. Soil Biol Biochem 32:1867–1875 Trasar-Cepeda C, Leiros MC, Gil-Sotres F (2008a) Hydrolytic enzyme activities in agricultural and forest soils. Some implications for their use as indicators of soil quality. Soil Biol Biochem 40:2146–2155 Trasar-Cepeda C, Leiros MC, Seoane S, Gil-Sotres F (2008b) Biochemical properties of soils under crop rotation. Appl Soil Ecol 39:133 Tu CM (1981) Effects of some pesticides on enzyme activities in an organic soil. Bulletin of Environmental Contamination 27:109–114 Tu CM (1993) Influence of ten herbicides on activities of microorganisms and enzymes in soils. Bulletin of Environmental Contamination and Toxicology 49:120–128 Visser S, Parkinson D (1992) Soil biological criteria as indicators of soil quality: soil microorganisms. Am J Altern Agric 7:33–37 Voet D, Voet JG (1995) Biochemistry. Introduction to enzymes. Chapter 12, 2nd edn. Wiley, New York, pp 332–344 Voets JP, Meerschman P, Verstraete W (1974) Soil microbiological and biochemical effects of long-term atrazine applications. Soil Biology and Biochemistry 6:149–152 Warkentin BP (1995) The changing concept of soil quality. J Soil Water Conserv 50:226–228 Yakovchenko VI, Sikora LJ, Rauffman DD (1996) A biologically based indicator of soil quality. Biol Fertil Soils 21:245–251 Yang L, Li T, Li F, Leucoff JH, Cohen S (2008) Fertilization regulates soil enzymatic activity and fertility dynamics in a cucumber field. Sci Hortic 116:21–26 Yeates GW, Orchard VA, Speir TW, Hunt JL, Hermans MCC (1994) Reduction in soil biological activity following pasture contamination by copper, chromium, arsenic timber preservative. Biology and Fertility of Soils 18:200–208 Yu YL, Shan M, Fang H, Wang X, Chu XQ (2006) Responses of soil microorganisms and enzymes to repeated applications of chlorothalonil. J Agric Food Chem 54:10070–10075 Zhang YL, Wang YS (2006) Soil enzyme activities with greenhouse subsurface irrigation. Pedosphere 16:512–518

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Zhang YM, Wu N, Zhou GY, Bao WK (2005) Changes in enzyme activities of spruce (Picea balfouriana) forest soil as related to burning in the eastern Qinghai-Tibetan plateau. Appl Soil Ecol 30:215–225 Zhang YL, Dai JL, Wang RQ, Zhang J (2008) Effects of long-term sewage irrigation on agricultural soil microbial structural and functional characterizations in Shandong, China. Eur J Soil Biol 44:84–91

Chapter 8

Enzyme Activities in the Rhizosphere of Plants Dilfuza Egamberdieva, Giancarlo Renella, Stephan Wirth, and Rafiq Islam

8.1

Introduction

The rhizosphere was first defined by Hiltner in 1904 as the soil volume surrounding the root surface. Generally, the rhizosphere has a thickness of 1–2 mm, but functionally, the rhizosphere can be defined as the soil portion physically and chemically influenced by growth and activity of the root. Therefore, the dimension of the rhizosphere can vary depending on the mass and architecture of the plant roots. The rhizosphere is a unique hot spot in soil from the viewpoint of microbial ecology, as soil micro-organisms are considerably stimulated by the activity of the roots (Jones et al. 2004; Hinsinger et al. 2006). Microbial communities are part of a complex food web that uses considerable amount of carbon fixed by the plant and released into the rhizosphere (i.e., rhizodeposits). In fact, increased microbial activity in the rhizosphere is sustained by nutrients secreted by plant roots in the form of soluble exudates, such as carbohydrates, aminoacids, low molecular weight organic acids and other photosynthates (Nannipieri et al. 2003). Generally, the rhizosphere soil contains more soluble sugars but less insoluble material than the surrounding soil and there is less nitrogen but more polyphenols (Somers et al. 2004; Raaijmakers et al. 2009). However, through the exudation of a wide variety of compounds, the plant roots may not only regulate positively or negatively the

D. Egamberdieva (*) Faculty of Biology and Soil Sciences, The National University of Uzbekistan, Tashkent 100174, Uzbekistan e-mail: [email protected] G. Renella Plant Nutrition and Soil Science, University of Florence, Florence, Italy S. Wirth Leibniz-Centre for Agricultural Landscape Research, Institute of Landscape Matter Dynamics, M€ uncheberg, Germany R. Islam Soil and Water Resources, Ohio State University South Centers, Piketon, OH, USA

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microbial activity in the rhizosphere but also regulate complex ecological interactions between plant and herbivores, plant beneficial microrganisms, and change the chemical and physical properties of the soil, and inhibit the growth of competing plant species (Nardi et al. 2000). Since plant nutrient uptake process occur through the rhizosphere, the activity of rhizosphere microbial community is of great importance for plant growth (Chanway 2002). The overall enzyme activity of the rhizosphere as well as bulk soil can depend on enzymes localized in root cells, root remains, microbial cells, microbial cell debris, microfaunal cells and the related cell debris, free extracellular enzymes or enzymes adsorbed onto or occluded into the soil colloids (Nannipieri et al. 2003). The study of different hydrolase enzyme activities in the rhizosphere soil and their changes is important since they indicate the potential of a soil to carry out specific biochemical reactions, and these hydrolytic enzymes are important in maintaining soil fertility and plant productivity (Burns 1982). Because plant nutrient uptake occurs through the rhizosphere, the activity of rhizosphere microbial community is of great importance for plant growth (Chanway 2002). Soil enzymes are involved in the catalysis of a large number of reactions necessary for life processes of microorganisms in soils, decomposition of organic residues, cycling of nutrients, and formation of organic matter and soil structure (Dick et al. 1996; Drijber et al. 2000; Colombo et al. 2002; Nannipieri et al. 2002). These enzymes include amylase, arylsulphatases, b-glucosidase, cellulase, chitinase, dehydrogenase, phosphatase, protease, urease and others, derived from plant, animal, or microbial origins (Dick and Tabatabai 1984; James et al. 1991; Gupta et al. 1994; Nannipieri et al. 2003). These enzymes can be accumulated, stabilized, and/or decomposed in the soil (Dick 1997). Plant roots have been considered a source of extracellular enzymes in soil. In early studies Estermann and McLaren (1961) found that root caps disruption of barley (Hordeum vulgare) possessed phosphatase activity. Juma and Tabatabai (1988) reported that corn (Zea mays) and soybean (Glycine max) roots contain acid phosphatase, but no detectable alkaline phosphatase activity. Enzyme activity in the rhizosphere can be of intracellular origin, released after microbial cell disuption or root cell sloughing (Hawes et al. 2003), and may be associated with soil colloids and cell debris, or enzymes may be actively secreted by plant roots or root associated microrganisms. Although the former enzyme location are supposed to contribute less to the whole rhizosphere enzyme activity being mineralized by microbial communities, it should be underlined that the methods today available for measuring the soil enzyme activity do not allow to discriminate the contribution of the different enzyme locations to the whole soil or rhizosphere enzyme activity (Nannipieri et al. 2003). Generally, low molecular organic compounds are passively released due to the existing steep intracellular/extracellular gradient whilst larger and more complex compounds or polymers are actively secreted through exocytosis (Neumann and R€ omheld 2007). It is important to take into account that the root exudation rate is higher in the root apical zone and decreases in the senescent root districts and that the root exudation profile is different in the various parts of the root.

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Rhizosphere microrganisms release extracellular enzymes for the initial degradation of high molecular polymers such as cellulose, chitin and lignin, proteins, leading to their mineralization to mineral N, P, and S (Burns 1982; Nannipieri et al. 1996). Badalucco et al. (1996) reported that in wheat rhizosphere various hydrolyse activites were higher in the rhizosphere than in the bulk soil with an increasing gradient towards the rhizoplane. Tarafdar and Jungk (1987) reported increasing of mineral P, decreasing organic P and progressive P depletion moving from bulk soil to the rhizoplane which was correlated to increasing phosphatase activity. For example, to our knowledge very few studies on the effects of the root mucilages on the enzyme stabilization and on the substrate diffusion have been conducted. Concerning the importance of enzyme diffusion, an interesting experiment was presented by Pilar et al (2009), showing that resorcinol-immobilized phosphatases increased the organic P fraction, enhanced the seed germination and had a positive effect on plant biomass and plant P content. Donegan and Seidler (1999) reported reduced dehydrogenase and alkaline phosphatase activities in the rhizosphere of transgenic alfalfa regardless of association with recombinant nitrogenfixing soil Sinorhizobium meliloti, as compared to rhizosphere soil sampled from parental alfalfa. George et al (2005) reported that phytase released by transgenic Trifolium subterraneum L. was inactivated by its interaction with soil colloids, or by proteolysis. Some information on the contribution of plant roots and microrganisms to the whole rhizosphere enzyme activity is provided by experiments based on transgenic plants, or based on rhizobacteria inocula. Richardson and Brinson (2001) reported that Arabidopsis thaliana containing a phytase gene (phyA) from Aspergillus niger absorbed P from various organic P sources, while George et al. (2005) reported that T. subterraneum L expressing a phytase gene (phyA) from Aspergillus niger released phytase. Lung et al. (2005) reported that Nicotiana tabacum carrying a phytase gene from Bacillus subtilis (168phyA), secreted extracellular phytase and used phytate as the sole P source. Concerning the contribution of rhizobacteria, information has been gained by studies involving the inoculation of genetically modified microrganisms. Naseby and Lynch (1997, 1998) were among the first to evaluate the effects of genetically modified rhizobatceria on the enzyme activities in the rhizosphere, and reported that inoculation of different Pseudomonas fluorescens strains had negative, neutral or positive effects on rhizosphere hydrolase activity. These findings may be attributable to the influence of other microbial groups, the availability of nutrients in the rhizosphere, plant species composition and its nutritional status, and also by the type of genetic modification (Naseby and Lynch 2002). Viterbo et al. (2002) reported the production of chitobiosidase, endochitinase, endo-b-1-3-glucanase and N-acetylglucosaminidase by the soil borne fungus Trichoderma harzianum Th008, soybean (Glycine max cv. Williams 82) roots, and with the exception of endochitinase. Furthermore, they found that T. Harzianum Th008 was the source of the endochitinase in the rhizosphere, whilst N-acetylglucosaminidase was mainly produced by the soybean roots. Idriss et al. (2002) reported that incoluation of Bacillus amyloliquefaciens FZB45 stimulated growth of maize growth under

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P limitating conditions in the presence of phytate, whereas a phytase-negative mutant strain (FZB45/M2) did not stimulate plant growth. However, in this case the experimental set-up did not exclude the contribution of plant phosphatases. Increasing production of phenol oxidase activities in sorghum root exudates of plants exposed to increased phenanthrene concentrations reported by Muratova et al. (2009) indicate an active role of enzymes released by root exudates, thus providing a positive plant response in the rhizosphere degradation of PAHs and derivatives in plants growing in contaminated soils. However, the importance of the production of intermediates in the degradation of aromatic compounds mediated by oxo-reductase enzyme activties illustrates once again the importance of the synergistic contribution of both plant and microbe derived enzymes in the rhizosphere (Sipil€a et al. 2008).

8.2

Effect of Root Exudates on Rhizosphere Soil Enzyme Activities

In the rhizosphere, up to 50% of the photosynthates are moved into roots, about 1% is actively released as root exudates but 10% is the total loss as root debris (Uren 2007). Root exudates represent a carbon-rich substrate for the rhizosphere microorganisms. Typical soluble root exudates are organic acids such as citrate, malate, fumarate, oxalate and acetate and carbohydrates such as glucose, xylose, fructose, maltose, sucrose, galactose, and ribose (Lugtenberg and Bloemberg 2004). Roots also release inorganic compound such as CO2, inorganic ions, protons and anions as a consequence of the root metabolic activity. Soluble organic root exudates are generally readily available to the rhizosphere and rhizoplane microorganisms and may diffuse at a longer distance from the rhizoplane than high-molecular weight compounds,which must be firstly hydrolysed in smaller compounds before they can be taken up by microbial cells (Nannipieri 2007). Root exudates can be a source of easily degradable N-compounds, such as amino acids and small peptides, able to induce protease synthesis (Garcia-Gil et al. 2004). Because of the high complexity of the rhizosphere environment, the mineralization of the different root exudates and their stimulatory effects on microbial activity have been approached, studying the effects of single low molecular weight organic molecules in simple systems mimicking the rhizosphere environment (Kozdro´j and van Elsas 2000; Badalucco and Kuikman 2001; Falchini et al. 2003; Baudoin et al. 2003). It is well established that N transformations in the rhizosphere soil are related to C dynamics and release of available C from roots (Qian et al. 1997). Gross N mineralization and immobilization rates of the rhizosphere soil are higher than those of the bulk soil, this is due to higher microbial activity in the rhizosphere than in the bulk soil. Landi et al. (2006) studied the effects of various model root exudates on N immobilization rates and reported that glucose was more effective

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than oxalic acid presumably because the first stimulated a larger proportion of soil micro organisms (Anderson and Domsch 1978), whereas the latter is decomposed by specialized microorganisms (Messini and Favilli 1990). In fact, fewer changes in the bacterial community induced during glucose decomposition compared to oxalic acid in model rhizosphere systems have been reported by Falchini et al. (2003), Baudoin et al. (2003), and Landi et al. (2006). In a follow up study, Renella et al (2007) reported that the effects of root exudates on N mineralization also depended on the soil chemical properties. Concerning the nitrogen dynamics in the rhizosphere, generally higher diversity of functional genes such as amoA and nifH genes have been detected in the rhizosphere than in bulk soil (Briones et al. 2003; Cocking 2003). Using the model rhizosphere system described by Badalucco and Kuikman (2001), Falchini et al. (2003) showed that selected root exudates were mineralized to different extents and had different stimulatory effects on microbial growth and on hydrolase activities, mostly localized in the rhizosphere zone. These studies have confirmed that root exudation is the main factor controlling microbial activity and community structure in the rhizosphere. The complex role of flavonoids released by white lupin roots in P mobilization from soil under P-deficiency conditions was studied by Tomasi et al. (2008) who showed that such polyphenolic compounds mobilize insoluble P, but also selectively inhibit the soil phosphohydrolase activity and reduce the microbial activity, possibly preventing the microbial P immobilization in the rhizosphere.

8.3

The Role of Lytic Enzymes and Plant Growth Regulators

Rhizosphere bacteria produce and release lytic enzymes that can hydrolyze a wide variety of polymeric compounds, including chitin, proteins, cellulose, and hemicellulose (Burns 1982; Lugtenberg and Bloemberg 2004; Pal and McSpadden Gardener 2006; Nannipieri 2007). The production of lytic enzymes by rhizosphere microorganisms can result in the suppression of plant pathogenic fungi directly. The rhizosphere bacterial species such Pseudomonas, Bacillus, Enterobacter, Alcaligenes, and Pantoea produced lytic enzymes such lipase, protease, collagenase, amylase, pectinase, cellulase, protease, lecithinase and were active against soil fungi (Lugtenberg et al. 2001; Egamberdiyeva and Hoflich 2003a, b, 2004; Egamberdieva et al. 2008). Pseudomonas is known to produce lytic enzymes and some isolates have been shown to be effective at suppressing fungal plant pathogens in tomatoes (Lugtenberg et al. 2001). Dunne et al. (2000) showed that overproduction of extracellular protease in the mutant strains of Stenotrophomonas maltophilia W81 resulted in improved biocontrol of Pythium ultimum. Nielsen and Sorensen (1997) demonstrated that isolates of P. fluorescens antagonistic to R. solani and P. ultimum, produced lytic enzymes. Plant growth promoting regulators such as auxin, or gibberellins are present in root exudates and thus enter the rhizosphere. Auxins are a class of plant hormones: the

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most common and well characterised is indol-3-acetic acid (IAA), which is known to stimulate both rapid (e. g., increases in cell elongation) and long term (e. g., cell division and differentiation) responses in plants (Cleland 1990; Hagen 1990). The auxin level was usually higher in the rhizosphere than in the free bulk soil, probably as a consequence of an increased microbial population or of accelerated metabolism owing to the presence of root exudates (Frankenberger and Arshad 1995; Somers et al. 2004). Microorganisms in the rhizosphere of various crops appear to have a greater potential to synthesize and release auxins as secondary metabolits because of the rich supply of substances and it is an important factor in soil fertility (Patten and Glick 2002; Frankenberger and Arshad 1995). According to several reports 86% of the bacterial isolates from the rhizosphere of various plants produced phytohormones such auxins, gibberelins, kinetin-like substances, but also different hydrolytic enzymes such protease, lipase, pectinase, and amylase (Hagen 1990; Frankenberger and Arshad 1995; Nielsen and Sorensen 1999; Lugtenberg et al. 2001). Lebuhn and Hartmann (1993) suggested the presence of higher auxin content in rhiosphere soils because of root colonization with Azospirilla and Rhizobium capable of excreting auxin without the addition of tryptophan (Patten and Glick 2002). Plant growth promoting rhizobacteria (PGPR) can affect plant growth directly by the synthesis of phytohormones, nitrogen fixation for plant use, improvement of nutrient uptake, solubilization of inorganic phosphate, and mineralization of organic phosphate (Dobbelaere et al. 2003). These physiological changes are linked to increases in enzyme activity. The exploitation of enzymes in the rhizosphere, weather or not extracellular, that can rapidly convert precursors to biologically active molecules, holds potential for increasing crop production (Zahir et al. 2001). In our study we observed that indole3-acetic acid (IAA) and gibberellic acid (GA) effect positively on alkaline phosphomonoestrase, phosphodiestrase, protease and urease activities in the model root surface (MRS) of a simplified rhizosphere system. Acid and alkaline phosphatase activities in wheat rhizosphere were strongly correlated with the depletion of organic P (Tarafdar and Jungk 1987). The plant growth regulators did not have an effect on acidic phosphomonoesterase activity in the rhizosphere (MRS) (Fig. 8.1). Protease activity is involved in the hydrolysis of N compounds to NH4, using low-molecular-weight protein substrates and microorganisms is responsible for breaking down urea into ammonium (Tabatabai 1982). Urease enzyme is responsible for the hydrolysis of urea fertiliser applied to the soil into NH3 and CO2 with the concomitant rise in soil pH (Byrnes and Amberger 1989). Other authors reported that urease released by plant root and rhizosphere micro organisms as intra- and extra-cellular enzymes (Burns 1986; Mobley and Hausinger 1989). Their activities in rhizosphere soil were strongly stimulated by IAA and GA (Fig 8.2). In early studies, Doyle and Stotzky (1993) found no difference in enzyme activities (arylsulfatase, phosphatases, and dehydrogenase) when an Escherichia coli strain was introduced into soil. Mawdsley and Burns (1994) reported that inoculating wheat seedling with Flavobacterium spp. increased activity of

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12000 water

IAA

GA

mg p-nitrophenol kg-1 so

10000

8000

6000

4000

2000

0 Phosphomonoestrase alcaline

Phosphomonoestrase acidic

Phosphodiestrase

Fig. 8.1 The effect of indol-3-acetic acid (IAA), and gibberellic acid (GA) on phosphomonoestrases and phosphodiesterase activities in rhizosphere soil (mg p-nitrophenol kg 1 soil h 1)

mg NH4+ –N kg soil

250

Protease

Urease

200 150 100 50 0 water

IAA

gibberelic acid

Fig. 8.2 The effect of indol-3-acetic acid (IAA), and gibberellic acid (GA) on protease and urease activities in rhizosphere soil (mg NH4+-N g 1 soil h 1)

OL-galactosidase, I-galactosidase, a-glucosidase, and I-glucosidase. This strain is able to produce plant growth stimulating substances such as auxin. Naseby and Lynch (1997) observed that bacterial inoculant P. fluorescens increased urease activity and decreased alkaline phosphomonoesterase and phosphodiesterase activity in the rhizosphere soil of wheat. This decrease explained by Naseby and

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Lynch (1997) that the inoculant inhabited niches of other microbes in the community would respond in a more dynamic fashion to the change in conditions. The first possibility is that the inoculant is competitively excluding certain microbial populations (De Leij et al. 1993). Other possible effects of the inoculant could be metabolic activity of the strain, that might have directly affected the indigenous microbial community (Keel et al. 1992) or modified root secretions (Mozafar et al. 1992). In earlier studies of plant growth regulators, the activities of rhizosphere bacteria including nitrogen fixation (Lindberg et al. 1985), production of cytokinin, auxin (Lebuhn et al. 1997; Timmusk et al. 1999), or hydrolytic enzymes such protease, lipase, pectinase, amylase (Nielsen and Sorensen, 1997) increased the N, P, and K uptake of plant components (Hoflich et al. 1997; Egamberdiyeva and Hoflich 2004). Tarafdar and Jungk (1987) observed that total P and organic P contents decreased in the rhizosphere soil of wheat, whereas the inorganic P content increased in the vicinity of the rhizoplane. Such an increase was correlated with the increased acid and alkaline phosphatase activities and fungal and bacterial population in the rhizosphere soil.

8.4

Effect of Soil Management and Heavy Metals on Rhizosphere Soil Enzyme Activities

Soil enzyme activities are considered to hold the potential to discriminate between soil management treatments, probably because they are closely related to microbial biomass, which is frequently reported to be sensitive indicator to such treatments (Dick 1997). Measurement of soil hydrolases provides an early indication of changes in soil fertility, since they are related to the mineralization of such important nutrients elements as N, P, C (Ceccanti and Garcia 1994). The crop management practices, and crop varieties affect the amount of acid phosphatase exuded by plant roots (Izaguirre-Mayoral et al. 2002; Wright and Reddy 2001). Yadav and Tarafdar (2001) reported that legumes secrete more phosphatase activity than cereal, and chickpea roots secrete more than maize (Li et al. 2004), which could be due to a higher requirement of P by legumes in the symbiotic nitrogen fixation process as compared to cereals. Other studies (Kai et al. 2002; Li et al. 2002) reported that P deficiency in the soil increased acid phosphotase secretion from plant root, which enhance the solubilisation and remobilisation of phosphate. It influences positively the ability of the plant to cope with P-stressed conditions. Urease enzyme also affected by soil type, temperature, organic matter content, soil management practices, crops and heavy metals (Yang et al. 2006). Gianfreda and Ruggiero (2006) reported that the enzyme activities in soil are affected by the presence and nature of the plant cover. Soils with a long history of continuous corn monoculture, without proper amendment with organic matter, showed low organic matter contents and low dehydrogenase, invertase,

Table 8.1 Acid, alcalic phosphomonoestrases, phosphodiestrase, galactosidase, glucosidase, urease, saline (MS) and strong saline (SS) soils Treatments (mg p-nitrophenol kg 1 soil  h 1) Acid phosphatase Alkaline phosphatase Phosphodiestrase Galactosidase EC NS 821.3 2,811.5 914.8 442.7 (1.3 dS m 1) 456.1 2,367.8 761.2 482.0 MS (5.6 dS m 1) 952.2 1,884.0 796.2 491.6 SS (7.1 dS m 1) 6.5 5.8

256.9 200.1 127.6

17.0

22.2

6.1

12.0

23.9

7.9

Glucosidase

23.9

(mg NH4+–N kg 1 soil  h 1) Urease Protease FDA

protease and FDA activities in non saline (NS), mid

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arylsulphatase, and b-glucosidase as compared with continuous corn-fertilized soils. A long-term intensive monoculture usually supplies lower amounts and diversity of organic matter than crop rotation, thus suppressing microbial activities ad consequently decreasing enzymatic ones (Klose and Tabatabai 2000). Soil enzyme activities are inhibited in trace element contaminated soils (Tyler et al. 1989) mainly due to direct interactions between trace elements with enzyme molecules, or substrates of enzyme–substrate complexes. In the rhizosphere the effects of trace elements on the enzyme activity are even more complex than in bulk soil due to the larger concentrations of trace element ligands in rhizodeposits, acidification and root trace element uptake. In spite of the relevance of the topic, there are relatively few studies on the hydrolase activity in the rhizospehere of trace element contaminated soils. Renella et al. (2005) reported a reduced hydrolase actvity in reposnse to the release of model root exudates by a model rhizosphere system in Cd contaminated soils. Possible causes of lower enzyme production in the rhizosphere of trace element contaminated soils could be both microbial metabolic stress (Renella et al. 2006) and lower mineralization of low molecular weight organic acids complexed with trace elements by soil microbial communities. To our knowledge, no information is available on the changes in the release of enzymes by plants exposed to trace element contaminated soils. More studies are surely needed to better understad the effects of trace element pollution on the hydrolase activity in the rhizosphere.

8.5

Impact of Salinity on Rhizosphere Enzyme Activities

Salinity is a major concern for irrigated agriculture in arid and semi-arid regions of the world (Shirokova et al. 2000; Egamberdiyeva et al. 2007). Eivazi and Tabatabai (1988), Garcı´a et al. (1994), and Batra and Manna (1997) showed that the activities of different soil enzymes were seriously reduced in saline soils affecting the capacity of the soil to recycle nutrients and to release them for their use by plants. Frankenberger and Bingham (1982) also reported that soil enzymes such dehydrogenase, phosphatase, sulfatase, amylase, and b-glucosidase activity was severely inhibited in salinized soils and their variation in soils seemed to be related to the physicochemical microbial properties of soils (Nannipieri et al. 1990; Zahir et al. 2001). Among the variables assessed, enzymatic activities (ureases, proteases, phosphatases, and glucosidases) may be sensitive indicators to detect changes occurring in soils under field conditions. The decline in enzymes activity with increasing salinity appeared to be associated with change in osmotic potential of the soil due to higher salt concentrations and specific ion toxicity (Zahir et al. 2001). Matsuguchi and Sakai (1995) investigated the effect of soil salinity with intensive cultivation on microbial communities in the soil-root system, and they found a negative effect of salt stress on microbial populations. Frankenberger and Bingham (1982) have pointed out the inhibition of different enzymatic activities caused by soil salinity. Dash and Panda (2001) established that NaCl salt stress induced changes in the

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growth and enzyme activities in blackgram (Phaseolus mungo) seeds, this negative effect was even greater on hydrolases (phosphatase and b-glucosidase). In spite of the large information on the effects of soil secondary salinization on soil enzyme activity, very few studies have focussed on the effects of salinization on the enzyme activity in the rhizosphere. In Table 8.1 we report rhizosphere soil enzyme activities in cotton grown on various levels of saline soil. We observed that salinity inhibited urease, protease, alkalic phosphomonoestrases and phosphodiestrase activity of rhizosphere soil. Non-saline soil showed highest alkaline phosphomonoestrase, phosphodiestrase, glucosidase, protease, urease, and fluorescein diacetate (FDA) (Table 8.1). However, alkaline phosphatase was higher in all soils than other hydrolase enzymes. Generally, alkaline phosphatase predominated in soils with neutral or slightly alkaline pH (Tripathi et al. 2007). Since, higher plants are devoid of alkaline phosphatase, the alkaline phosphatase of soils seems to be derived totally from microorganisms (Dick and Tabatabai 1983; Juma and Tabatabai 1988). Gianfreda and Bollag (1994) observed a direct inhibitory effect of soil organic constituents on the activities of acid phosphatase. Acid phosphomonoestrase, phosphodiestrase, and galactosidase were not affected by an increase of soil salinity, and did not correlate with soil electrical conductivity, organic matter content, Cl, and Na (Table 8.1). Other enzymes such alkaline phosphomonoestrases, proteases, glucosidases, ureases, and fluorescein diacetate hydrolase were inhibited in saline soils, in relation to the soil organic C and microbial biomass (Table 8.1). Carrasco et al. (2006) found that the changes in microbiological activity along the spatial vegetation gradient were also revealed by the variations in protease-BAA, phosphatase, urease and b-glucosidase activities. Urease and protease activities appear to be more sensitive to salinity than phosphatases. Garcı´a et al. (1994) reported that the reduction in enzyme activities can be due to lower microbial biomass that releasing less enzymes but also by the fact that in semi-arid soils the enzyme activity is mainly extracellular, stable and form complexes with the organic and mineral colloids. In saline soils, salt tolerant bacteria produce enzymes, whose activity has a greater salt requirement than that of corresponding enzymes produced by non- salt tolerant bacteria (Zahran 1997). This decrease in enzymatic activity with soil salinity can be explained by the fact that in the semi arid soils many of the enzymes are extracellular, stable and form complexes with the organic and mineral colloids (Garcı´a et al. 1994; Garcia and Hernandez 1996). The increase of conductivity disperses the clays and the stable enzymes remain unprotected and therefore, more susceptible to denaturation (Frankenberger and Bingham 1982; Pathak and Rao 1998). Garcia and Hernandez (1996) in their work reported that salinity negatively affects biological and biochemical fertility of the soils and is more pronounced with NaCl than Na2SO4 which can be attributed to the toxic effect of a particular ion in saline soils on microbial growth. Probably toxicity of chlorides was greater than the toxicity by sulphates with those enzymes and microbial biomass being the most sensitive. In addition, high salt concentrations tend to reduce the solubility and denature enzyme proteins through disruption of the tertiary protein structure which is essential for enzymatic activity (Frankenberger and Bingham 1982;

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Zahran 1997). Reboreda and Cac¸ador (2008) reported that extracellular enzyme activity in salt marshes sediments colonised by Spartina maritima were positively correlated with root biomass indicating the role of halophyte roots on nutrient cycling and microbial functioning sediments.

8.6

Conclusions

Research on the enzyme activity in the rhizosphere suffer major methodological limitations due to the complexity of the rhizosphere in terms of sampling, and due to the susceptibility of the rhizosphere to artifacts during rhizosphere sampling. For such reasons, enzyme activity in the rhizosphere has been studied by means of artificial systems mimicking the release of selected root exudates under controlled conditions. Although this approach is useful to describe specific effects, we all are aware that they are very far from the real plant soil microbe situation. A possible approach is the direct visualization of the rhizosphere environment. The use of soil thin sections, treated for ultracytochemical localization of enzymes in rhizobacteria and root extracellular polysaccharides and cell wall by transmission electron microscopy were proposed by R.C. Foster (see Ladd et al. 1996), but these approaches were not sufficiently developed and can not be applied for enzymes associated to electron-dense particles and minerals. Concerning the relationship between microbial diversity and enzyme activity in the rhizosphere, in spite of the increasing knowledge on the microbial community composition, no comparisons with the enzyme activity are normally carried out. This kind of comparitve work may be of great interest from theoretical and practical aspects. For example, it may clarify whether microbial inocula are capable of increasing specific enzyme activities for various purposes, such as reducing the crop fertilization, or enhancing the phytodegradation of organic pollutants. Finally, a transition from studies determining only the enzyme activity to studies aiming at identifyng and quantifying the enzyme molecules by the proteomic approach is in our opinion, a better approach for understanding of the source and relevance of hydrolase activity in the rhizosphere. For example, Wen et al. (2007) showed that enzymes released in the pea root secretome may play a role in whole protection mechanisms in the presence of pathogen fungi. Identification of these enzymes may be useful to improve the crop protection strategy. De la Pen˜a et al. (2008) studied the role of proteins secreted by secreted roots of Medicago sativa and Arabidopsis thaliana in the presence of symbiotic (Sinorhizobium meliloti) and pathogenic (Pseudomonas syringae) bacteria, and found that the interaction between the two plants increased the secretion of various proteins, prevalently hydrolases, peptidases, as compared to control plants. Implementation of rhizosphere proteomics for disentangling the complexity of the enzyme secretion by plants and microrganisms in the plant-microbial chemical communication in the rhizosphere may likely improve our knowledge on the enzyme activity in the rhizosphere more than other technical improvements. It is considered a key issue for elucidating the importance and the location of the enzyme activity in the

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rhizosphere in the next few years from both applicative and basic research viewpoints.

References Anderson JPE, Domsch KH (1978) A physiological method for the